The RNA-DNA Nexus: Unveiling the Molecular Machinery of Life, and the Intelligent Design Paradigm

6. Decarboxylation to form UMP

The final reaction in the pathway is the decarboxylation of orotidine-5'-monophosphate (OMP) by the enzyme OMP decarboxylase (ODCase), resulting in the formation of uridine-5'-monophosphate (UMP). This reaction is crucial for the biosynthesis of pyrimidine nucleotides, which are essential building blocks for RNA and DNA.

OMP decarboxylase

OMP decarboxylase is an exceptional enzyme in terms of catalytic efficiency. It is known for being an extraordinarily efficient catalyst capable of accelerating the uncatalyzed reaction rate by a factor of 10^17. To put this in perspective, the uncatalyzed reaction which would take 78 million years to convert half the reactants into products is accelerated to 18 milliseconds when catalyzed by OMP decarboxylase. It enhances the rate of the decarboxylation reaction by an astonishing factor of 2 × 10^23 compared to the uncatalyzed reaction. This makes it one of the most catalytically proficient enzymes known in the biological world. To put the magnitude of the rate enhancement into perspective, let's consider a hypothetical scenario. Suppose the uncatalyzed reaction without ODCase has a rate constant (k_uncat) of 1 per second. With ODCase's catalytic efficiency, the rate constant (k_cat) of the reaction would be 2 × 10^23 per second. The catalyzed reaction occurs 2 × 10^23 times faster than the uncatalyzed reaction.

If the catalyzed reaction were to take 1 second, the uncatalyzed reaction would be taking approximately 200 billion years.



Interestingly, the catalytic activity of ODCase does not rely on any cofactors or metal ions to stabilize the putative carbanion intermediate formed during the reaction. The catalysis relies on a handful of charged amino acid residues positioned within the active site of the enzyme. Instead, the enzyme achieves its remarkable catalytic proficiency through precise molecular interactions and binding energy. The exact mechanism by which ODCase catalyzes the decarboxylation reaction is not fully understood. However, it is known that the removal of the phosphate group from OMP, which is located far away from the carboxyl group at C6, significantly decreases the reaction rate by a factor of 7 × 10^7. This phenomenon highlights the concept of preferential transition state binding. Preferential transition state binding refers to the enzyme's ability to selectively bind and stabilize the transition state of a reaction over the substrate or product. In the case of ODCase, the enzyme's active site is optimized to bind the transition state of the decarboxylation reaction more favorably than the substrate OMP or the product UMP. This preferential binding of the transition state leads to a significant enhancement in the reaction rate.


Homodimer Orotidine 5′-monophosphate decarboxylase (ODCase)   One subunit is colored red and the other blue. Completely conserved residues are emphasized with ball-and-stick representations. The inhibitor, BMP, is drawn in yellow. (A) Viewed through the R/â-barrel. (B) Viewed perpendicular to the view in panel A. . The two subunits are connected by a series of hydrogen bonds between residues His24 and Asp90; Lys73 and Asp76; N79 and Asp200; Arg105 and Ser132, Asp137, and Asp200; His75 and Lys47 and His99; and Asp200 and N79.


A Proficient Enzyme




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Reaction scheme of OMP decarboxylation.

Enzyme Catalyst: Enzyme expert Dr Richard Wolfenden, of the University of North Carolina, showed in 1998 that a reaction ‘“absolutely essential” in creating the building blocks of DNA and RNA would take 78 million years in water’, but was speeded up 10^18 times by an enzyme.1 This was orotidine 5′-monophosphate decarboxylase, responsible for de novo synthesis of uridine 5′-phosphate, an essential precursor of RNA and DNA, by decarboxylating orotidine 5′-monophosphate (OMP). In 2003, Wolfenden found another enzyme exceeded even this vast rate enhancement. A phosphatase, which catalyzes the hydrolysis of phosphate dianions, magnified the reaction rate by thousand times more than even that previous enzyme—10^21 times. That is, the phosphatase allows reactions vital for cell signalling and regulation to take place in a hundredth of a second. Without the enzyme, this essential reaction would take a trillion years—almost a hundred times even the supposed evolutionary age of the universe (about 15 billion years)! Implications: Wolfenden said, ‘Without catalysts, there would be no life at all, from microbes to humans. It makes you wonder how natural selection operated in such a way as to produce a protein that got off the ground as a primitive catalyst for such an extraordinarily slow reaction.’

 CAD, the trifunctional protein

UMP (uridine monophosphate) biosynthesis, just described,  involves a series of enzymatic reactions that convert simple precursors into UMP. In bacteria, each of the six enzymes involved in UMP biosynthesis exists as independent proteins, meaning that they are encoded by separate genes and synthesized as separate polypeptide chains. However, in animals, the first three enzymatic activities of the UMP biosynthesis pathway occur on a single polypeptide chain. This means that a single large protein carries out the functions of three enzymes: carbamoyl phosphate synthetase II (CPS II), ATCase (aspartate transcarbamoylase), and dihydroorotase.

The 3-in-1 protein, or trifunctional protein,  that combines the activities of the first three proteins in the pathway is known as CAD (carbamoyl phosphate synthetase/aspartate transcarbamoylase/dihydroorotase). The CAD protein is a multifunctional enzyme that catalyzes the first three steps of the UMP biosynthesis pathway in a coordinated manner.

The CAD protein has a molecular weight of 243 kDa. This value indicates that CAD is a large-sized protein. Proteins can generally be categorized into small (< 20 kDa), medium (20-70 kDa), or large (> 70 kDa) based on their molecular weight. With a molecular weight of 243 kDa, the CAD protein falls into the large protein category. The CAD protein can be put into perspective with the size of the ribosome. The ribosome is significantly larger than the CAD protein. The combined size of the bacterial ribosome is approximately 2.5-3 million daltons (MDa), which is equivalent to 2,500-3,000 kDa. This makes the ribosome roughly 10-12 times larger than the CAD protein. The combined size of the eukaryotic ribosome is approximately 4.2-4.5 million daltons (MDa), which is equivalent to 4,200-4,500 kDa. Thus, the eukaryotic ribosome is about 17-18 times larger than the CAD protein.

The active site of CAD is characterized by a carboxylated lysine residue, which acts as a bridge for two zinc ions with a positive charge (+2). Another zinc ion contributes to stabilizing a histidinate ion. These zinc ions and the lysine residue play crucial roles in the enzyme's activity.  The function of CAD is regulated by various molecules to modulate its enzymatic activity. The CAD protein forms a complex with multiple enzymatic activities to facilitate the efficient synthesis of pyrimidines in animals. There are several reasons why these activities are combined into a single protein complex:  By bringing together the enzymatic activities involved in different steps of pyrimidine synthesis, the CAD complex allows for tight coordination and efficient transfer of intermediates. The proximity of the active sites within the complex enables rapid channeling of substrates and intermediates between the different enzymatic reactions, minimizing losses and improving overall efficiency. The CAD complex enables the direct and efficient transfer of intermediates between the enzymatic activities without their release into the cellular environment. This substrate channeling helps to prevent the loss of intermediates and ensures their efficient utilization within the pathway, reducing wastage and enhancing the productivity of pyrimidine synthesis. The complex formation of CAD allows for coordinated regulation of the enzymatic activities. Regulatory mechanisms can act on the complex as a whole, affecting the overall activity of pyrimidine synthesis. Feedback inhibition by end products, such as UTP and UMP, can modulate the activity of the CAD complex, ensuring that the synthesis of pyrimidines is balanced according to cellular needs.  The assembly of the CAD complex provides structural stability to the individual enzymatic components. Some of the domains within CAD may not be stable on their own and require the presence of other domains in the complex for proper folding and stability. The complex formation ensures the integrity and functionality of each enzymatic activity within CAD.  The absence of the CAD complex in prokaryotes is likely a consequence of differences in genetic organization, metabolic flexibility between prokaryotes and eukaryotes. Prokaryotes have  alternative mechanisms to synthesize pyrimidines that are better suited to their specific needs and environmental conditions.

Pyrimidine Nucleotide Biosynthesis Is Regulated at ATCase or Carbamoyl Phosphate Synthetase II

The regulation of pyrimidine nucleotide biosynthesis differs between bacteria and animals, with bacteria primarily regulating the pathway at the aspartate transcarbamoylase (ATCase) reaction, while animals primarily regulate it at the carbamoyl phosphate synthetase II (CPS II) reaction.



In the regulation of pyrimidine biosynthesis, control networks play a crucial role in maintaining the balance of nucleotide production. These control networks differ between E. coli (representing bacteria) and animals, reflecting the specific regulatory mechanisms in each system.

(a) In E. coli: In E. coli, the control network for pyrimidine biosynthesis involves several key control points. The major control point is at the aspartate transcarbamoylase (ATCase) reaction, which is depicted by a red octagon. ATCase is allosterically stimulated by ATP (shown by a dashed green arrow), which enhances its catalytic activity. Conversely, CTP acts as a feedback inhibitor of ATCase (indicated by a dashed red arrow), reducing its activity and slowing down the production of pyrimidine nucleotides. Additionally, UTP, another pyrimidine nucleotide, can also inhibit ATCase in many bacterial species.

(b) In animals: In animals, the control network for pyrimidine biosynthesis differs from that in bacteria. The major control point in animals is at the carbamoyl phosphate synthetase II (CPS II) reaction, represented by a green circle. CPS II is activated by ATP and PRPP (5-phosphoribosyl-α-pyrophosphate) through allosteric regulation (dashed green arrows). The binding of ATP and PRPP enhances the activity of CPS II, promoting the synthesis of carbamoyl phosphate, a key intermediate in pyrimidine biosynthesis. On the other hand, UDP and UTP act as inhibitors of CPS II, reducing its activity and controlling the rate of pyrimidine nucleotide production (dashed red arrows).

The control networks depicted in both E. coli and animals illustrate the importance of feedback inhibition and activation in maintaining the appropriate levels of pyrimidine nucleotides. These regulatory mechanisms ensure that pyrimidine biosynthesis is tightly regulated and adapted to the metabolic needs of the organism. By responding to the levels of ATP, CTP, UTP, UDP, and PRPP, the control networks help maintain cellular homeostasis and prevent the excessive accumulation or depletion of pyrimidine nucleotides.

In bacteria, such as E. coli, the regulation of pyrimidine biosynthesis occurs mainly at the ATCase reaction. ATCase is a key enzyme that catalyzes the conversion of carbamoyl phosphate and aspartate into N-carbamoyl-L-aspartate. In E. coli, ATCase is regulated by allosteric mechanisms. It is allosterically stimulated by ATP and inhibited by CTP, which helps control the rate of pyrimidine biosynthesis. Additionally, in many bacterial species, UTP acts as a major inhibitor of ATCase. These allosteric regulations ensure that the production of pyrimidine nucleotides is balanced in response to the intracellular levels of ATP, CTP, and UTP. In animals, the regulation of pyrimidine biosynthesis is primarily governed by the activity of carbamoyl phosphate synthetase II (CPS II). CPS II is responsible for catalyzing the conversion of glutamine, ATP, and bicarbonate into carbamoyl phosphate, a crucial intermediate in pyrimidine biosynthesis. In animals, CPS II is regulated by feedback inhibition. UDP and UTP act as inhibitors of CPS II, limiting its activity and thus controlling the rate of pyrimidine nucleotide synthesis. On the other hand, ATP and PRPP (5-phosphoribosyl-α-pyrophosphate) act as activators of CPS II, stimulating its activity and promoting pyrimidine biosynthesis. In addition to ATCase and CPS II, a second level of control in the mammalian pathway occurs at OMP (orotidine 5'-monophosphate) decarboxylase. UMP and to a lesser extent CMP competitively inhibit OMP decarboxylase. This regulation ensures that the production of OMP, an intermediate in pyrimidine biosynthesis, is finely tuned according to the levels of UMP and CMP. The rate of OMP production in all organisms depends on the availability of its precursor, PRPP. The intracellular level of PRPP is influenced by the activity of ribose phosphate pyrophosphokinase, an enzyme that catalyzes the conversion of ribose 5-phosphate and ATP into PRPP. The activity of ribose phosphate pyrophosphokinase is inhibited by ADP and GDP, which provides another level of regulation in controlling the production of PRPP and, consequently, the overall rate of pyrimidine nucleotide biosynthesis.

The differences in the regulatory networks for pyrimidine biosynthesis between bacteria and eukaryotes are evidence that one network cannot evolve from the other.  The regulatory networks for pyrimidine biosynthesis in bacteria and eukaryotes exhibit distinct features and complexities that are finely tuned to the specific needs of each organism. These networks involve multiple enzymes, feedback mechanisms, and allosteric regulation, which require precise coordination to ensure the proper production of pyrimidine nucleotides. If one regulatory network were to evolve from the other, it would require numerous coordinated changes in enzyme function, gene expression, and regulatory interactions. The chance of these changes occurring simultaneously, while still maintaining a functional and efficient system, is highly improbable. Furthermore, the differences in the nitrogen source for amination (glutamine in animals and ammonia in bacteria) and the involvement of additional regulators (such as ATP and PRPP in eukaryotes) further highlight the distinct design and adaptation of the regulatory networks in each organism.  The intricate coordination and interdependence of these networks are indicative of purposeful design rather than random evolutionary processes.

UMP Is Converted to UTP and CTP

The conversion of UMP (uridine monophosphate) to UTP (uridine triphosphate) and CTP (cytidine triphosphate) is an essential step in nucleotide biosynthesis. This process involves enzymatic reactions that occur in both animals and bacteria, although there are some differences in the source of the amino group involved. In the synthesis of UTP, UMP is first phosphorylated to form UDP (uridine diphosphate) by the action of a nucleoside monophosphate kinase. This enzyme transfers a phosphate group from ATP (adenosine triphosphate) to UMP, resulting in the formation of UDP. Subsequently, UDP undergoes another phosphorylation reaction catalyzed by a nucleoside diphosphate kinase, which transfers a phosphate group from ATP to UDP, yielding UTP.

The formation of CTP from UTP involves the process of [url=https://en.wikipedia.org/wiki/Amination#:~:text=Amination is the process by,because organonitrogen compounds are pervasive.]amination[/url]. In animals, the amination of UTP to form CTP is facilitated by the enzyme CTP synthetase. During this reaction, the amino group required for the amination of UTP is donated by glutamine. Glutamine is an amino acid that serves as the nitrogen source in the synthesis of CTP in animals. The reaction catalyzed by CTP synthetase results in the replacement of the oxygen atom in the uracil ring of UTP with an amino group, leading to the formation of CTP. In bacteria, the process of CTP synthesis is slightly different. Instead of using glutamine as the amino group donor, bacteria directly utilize ammonia for amination. The enzyme responsible for the amination of UTP in bacteria is also called CTP synthetase, but it utilizes ammonia as the nitrogen source. This difference in the nitrogen source between animals and bacteria reflects their distinct metabolic pathways and adaptations to different environmental conditions.



The difference in the nitrogen source and the specific enzymatic mechanisms between animals and bacteria reflect their distinct metabolic pathways and adaptations to different environmental conditions. Glutamine is a readily available nitrogen source in animals, and the use of this amino acid as a nitrogen donor in CTP synthesis allows for efficient utilization of available nitrogen resources. On the other hand, bacteria can directly assimilate ammonia from their surroundings, making it a convenient nitrogen source for CTP synthesis.  In bacteria, the sole enzyme involved is CTP synthetase, which directly utilizes ammonia as the nitrogen source. In animals, two enzymes are involved: nucleoside diphosphate kinase, which converts UTP to UDP, and CTP synthetase, which utilizes glutamine as the nitrogen donor for the amination of UDP to form CTP.

Nucleotide metabolism: By evolution?

G. Caetano-Anollés (2013): The origin of metabolism has been linked to abiotic chemistries that existed in our planet at the beginning of life. While plausible chemical pathways have been proposed, including the synthesis of nucleobases, ribose and ribonucleotides, the cooption of these reactions by modern enzymes remains shrouded in mystery. Pathways of nucleotide biosynthesis, catabolism, and salvage originated ∼300 million years later by concerted enzymatic recruitments and gradual replacement of abiotic chemistries. The simultaneous appearance of purine biosynthesis and the ribosome probably fulfilled the expanding matter-energy and processing needs of genomic information. 1

Comment: These are assertions, clearly not based on scientific data and observations, but ad-hoc conclusions that lack evidence. 

Pyrimidine Bases can be salvaged and recycled

M.Lieberman (2017): Pyrimidine bases are normally salvaged by a two-step route. First, a relatively nonspecific pyrimidine nucleoside phosphorylase converts the pyrimidine bases to their respective nucleosides. Notice that the preferred direction for this reaction is the reverse phosphorylase reaction, in which phosphate is released and is not being used as a nucleophile to release the pyrimidine base from the nucleoside. The more specific nucleoside kinases then react with the nucleosides, forming nucleotides. As with purines, further phosphorylation is carried out by increasingly more specific kinases. The nucleoside phosphorylase–nucleoside kinase route for synthesis of pyrimidine nucleoside monophosphates is relatively inefficient for salvage of pyrimidine bases because of the very low concentration of the bases in plasma and tissues. 


1. Gustavo Caetano-Anollés: Structural Phylogenomics Reveals Gradual Evolutionary Replacement of Abiotic Chemistries by Protein Enzymes in Purine Metabolism March 13, 2013

5





Formation of Deoxyribonucleotides (DNA)

DNA is “the Blueprint of Life.” It contains the data needed to make every single protein that life can't go on without. No DNA, no proteins, no life. RNA has a limited coding capacity because it is unstable. A lot has been written and said about the fact that DNA, besides epigenetic information, stores the instructions to make every living organism on the planet. It is the blueprint of life. Less known or widespread is the question of the origin of the DNA molecule. RNA is a prominent molecule, in special in Origin of Life questions, popularized through the so-called RNA world. But what is the origin of the DNA molecule? As we will see, the making of DNA, starting from RNA, is an exceedingly complex process and requires some of the most complex proteins known, like Ribonucleotide reductase, or in short RNR enzymes, that come in three versions. I would say, everything equal, just by means of the complexity of RNR enzymes, which are vital for all life, abiogenesis is a failure. The enzyme is extraordinarily sophisticated, complex, and energy dispendious to have originated by natural means.

Why is RNA replaced by DNA? 



Donald Voet et.al. (2016): DNA differs chemically from RNA in two major respects: (1) Its nucleotides contain 2′-deoxyribose residues rather than ribose residues, and (2) it contains the base thymine (5-methyluracil) rather than uracil.

DNA without the code reading cell machinery can do nothing on its own, which is why the vital flame of life must be passed down from living cell to living cell, uninterrupted since the very beginning of life itself. The genetic program is sophisticated enough that it causes genes to be transcribed that produce proteins that are themselves transcription factors secreted out of the cell to instruct neighboring cells as to which of their genetic programs to begin running. It is this complex coordination, leading to the switching on or off of particular genes in other cells, that starts the process of building a whole multicellular organism. In this way, it is not just the genetic program that is necessary for building an animal, person, or plant, but the local chemical environment that the program of each cell finds itself living in. The chemical neighborhood is just as important as the genetic constituency.



Where two distinct processes are required to work together to perform a function, and individually, they do perform no function, then it is necessary for both components to arise together. Keeping a pool of functional DNA molecules depends on 1. The proper biosynthesis of the RNA and DNA molecules itself and 2. Kinga Nyíri (2018): The fine-tuned regulation of nucleotide metabolism to ensure DNA replication with high fidelity which is essential for proper development in all free-living organisms 1 One process depends on the other, and singularly, they would not convey what all organisms need.

Why does Thymine replace Uracil in DNA?

M.Cobb (2015): The replacement of RNA as the repository of genetic information by its more stable cousin, DNA, provides a more reliable way of transmitting information. This explains why DNA uses thymine (T) as one of its four informational bases, whereas RNA uses uracil (U) in its place. The problem is that cytosine (C), one of the two other bases, can easily turn into U, through a simple reaction called deamination. This takes place spontaneously dozens of times a day in each of your cells but is easily corrected by cellular machinery because, in DNA, U is meaningless. However, in RNA such a change would be significant – the cell would not be able to tell the difference between a U that was supposed to be there and needed to be acted upon, and a U that was a spontaneous mutation from C and needed to be corrected. This does not cause your cells any difficulty, because most RNA is so transient that it does not have time to mutate – in the case of messenger RNA it is copied from DNA immediately before being used. Thymine is much more stable and does not spontaneously change so easily. The adoption of DNA as the genetic material, with its built-in error-correction mechanism in the shape of the two complementary strands in the double helix, and the use of thymine in the sequence, provided a more reliable information store and slowed the rate of potentially damaging mutations.4

V. Thakur (2018): The only structural difference between Thymine and Uracil is the presence of methyl group in Thymine. This methyl group facilitates the repair of damaged DNA, providing an additional selective advantage. Cytosine in DNA undergoes spontaneous deamination at a perceptible rate to form Uracil. For example, under typical cellular conditions, deamination of Cytosine to Uracil (in DNA) occurs in about every 107  Cytidine residues in 24 hours, which means 100 spontaneous events per day. The deamination of Cytosine is potentially mutagenic because Uracil pairs with Adenine and this would lead to a decrease in G≡C base pairs and increase in A=U base pairs in DNA of all cells. Over the time period, the Cytosine deamination could completely eliminate G≡C base pairs. But, this mutation is prevented by a repair system that recognizes Uracil as foreign in DNA and removes it. Thus, the methyl group on thymine is a tag that distinguishes thymine from deaminated cytosine. But, if DNA normally contains Uracil recognition would be more difficult and unpaired Uracil would lead to permanent sequence changes as they were paired with Adenine during replication. So, we can say that Thymine is used in place of Uracil in DNA to enhance the fidelity of the genetic message. In contrast, RNA is not repaired and so Uracil is used in RNA because it is a less expensive building block.4

M.Eberlin (2019): Where DNA uses thymine (T) as one of its bases, RNA uses uracil (U). This U-to-T exchange is intriguing because the chemical structures of T and U are nearly identical, distinguished only by a single, small methyl group (CH3 ). As the editors of the NSTA WebNews Digest noted, converting uracil to thymine requires energy, so why do cells bother to methylate uracil into thymine for DNA? Additionally, the extra group is placed in what seems to be a rather inert position on the T ring. It seems therefore that such a rather small and inert CH3 group is there only to “differentiate” U and T while disturbing the chemical properties as little as possible. A number of evolutionary explanations have been offered for this U-to-T exchange, but it turns out this exchange maintains the integrity of the whole information storage system, so a fully evolved form of it would have been needed from the start. As we saw earlier, the four RNA bases—A, U, G, and C—are superb for the job they have, but they also cause a problem if used in the wrong context. The U-to-T exchange is the solution. The original quartet is fine for less stable RNA, but not the best choice for long-lasting DNA. The U base would still establish a preferential pairing with A, but the A=U pair is not ideal for the role DNA fills, since U can also match efficiently with all the other bases, including itself. DNA’s T, on the other hand, is much more selective than U in its pairing with adenine (A), forming a more stable A=T pair. This specificity makes sense when you remember that DNA, which is made of nucleic acids, phosphate anions, and sugar molecules, is very hydrophilic (water-loving). As Michael Onken explains, the addition of a hydrophobic CH3 group to U (thus forming T) causes T to repel the rest of the DNA. This, in turn, shifts T to a specific location in the helix. This perfect positioning causes T to bind exclusively with A, making DNA a better, more accurate information replication system. This guarantees the long-lasting integrity of DNA information. So we see that the most fundamental design principles of the DNA helix are carefully tuned for the code to work properly, from the number of H-bonds between the A=T and G≡C interactions to the exact fit of the molecules between the two wires that form the double helix.  6


Why has the oxygen-hydrogen (OH) group in RNA been replaced by hydrogen (H) in DNA? 

Gerald F. Joyce (2002): The primary advantage of DNA over RNA as a genetic material is the greater chemical stability of DNA, allowing much larger genomes based on DNA. Protein synthesis may require more genetic information than can be maintained by RNA. To expand, RNA is unsuitable for large genomes because the 2'-OH of ribose (obviously absent from the 2'-dexoyribose of DNA) renders the phosphodiester bond susceptible to alkaline hydrolysis. (Wikipedia: hydrolysis is a reaction in which a phosphodiester bond in the sugar-phosphate backbone of RNA is broken, cleaving the RNA molecule.)3

Jayachandran (2014): DNA is such an important molecule so it must be protected from decomposition and further reactions. The absence of one Oxygen is the key to extending DNA's longevity. When the 2' Oxygen is absent in deoxyribose, the sugar molecule is less likely to get involved in chemical reactions( the aggressive nature of Oxygen in chemical reactions is famous). So by removing the Oxygen from the deoxyribose molecule, DNA avoids being broken down. From an RNA point of view, Oxygen is helpful, unlike DNA, RNA is a short-term tool used by the cell to send messages and manufacture proteins as a part of gene expression. Simply speaking mRNA (Messenger RNA) has the duties of turning genes ON and OFF, when a gene needed to be put ON mRNA is made and to keep it OFF the mRNA is removed. So the OH group in 2' is used to decompose the RNA quickly thereby making those affected genes in OFF state.

M.Eberlin (2019): DNA must be highly stable, while RNA, as the temporary intermediate between DNA and protein must be dramatically less stable. RNA uses the intact ribose sugar molecule to make its polymeric wire, while DNA uses a de-oxygenated version of it—deoxyribose. Since an OH group has been replaced by an H at an apparently “chemically silent” 2’-position in the ribose ring, it seems strange at first sight to note such care for a seemingly trivial molecular detail. But it turns out that there is a crucial-for-life reason for this amazing chemical trick. The choice of D-ribose for m-RNA and D-deoxyribose for DNA increases the chemical stability of DNA while decreasing that of RNA in an alkaline medium. Both of these are for a reason. If nuclear DNA is the hard drive of life, storing information for the long term, messenger RNA (mRNA) is life’s flash drive, transmitting information over short periods of time. RNA’s lifetime had therefore to be short, otherwise, protein production would never stop. Life needed a way to quickly “digest” via hydrolysis and ideally recycle the components of RNA when its job is finished. When chemists analyzed this “mysterious” OH/H exchange, they discovered that the apparently “silent” 2’-OH group helps RNA undergo hydrolysis about one hundred times faster than DNA. So we see that ribose had to be used in RNA for easy digestion in an alkaline medium, and deoxyribose had to be used in DNA for longevity. Otherwise, life would be impossible. Again, by all appearances, this stability control for both DNA and RNA had to be anticipated ahead of time and the solution provided with just-in-time delivery.6

Ribonucleotide Reductase 

All cellular organisms have double-stranded DNA genomes. 26 There are no known life forms that do not use DNA as their genetic material. There are no known life forms that use other types of genetic material. Ribonucleotide reductase (RNR) is an enzyme that is essential for DNA synthesis in all cells. Logically, it follows, that it was present when life started, and therefore, its origin cannot be explained by evolutionary mechanisms. If a cell were to completely lack ribonucleotide reductase (RNR) enzymes, it would not be able to convert ribonucleotides (the building blocks of RNA) into deoxyribonucleotides (the building blocks of DNA). This would lead to a shortage of deoxyribonucleotides and an inability to synthesize DNA, resulting in severe disruption of DNA replication and repair. Without functional RNR enzymes, the cell would not be able to maintain its genome or carry out essential cellular functions that require DNA synthesis. This would lead to genomic instability, increased susceptibility to DNA damage, and eventual cell death.

Overview: 

Ribonucleotide reductase (RNR) enzymes are among the most sophisticated and complex enzymes known. They are complex multi-subunit enzymes that require a range of cofactors and allosteric regulators to function properly. They also undergo complex regulatory mechanisms, such as transcriptional, post-transcriptional, translational, and post-translational control, to maintain the appropriate balance of deoxyribonucleotides in the cell. Moreover, RNR enzymes have to be highly regulated and tightly controlled, with a variety of feedback mechanisms and checkpoints that ensure that the appropriate balance of deoxyribonucleotides is maintained in the cell. This complex regulation is necessary to prevent excessive or insufficient levels of deoxyribonucleotides, which can have serious consequences for DNA synthesis and repair, and ultimately for the survival and proliferation of the organism. RNR enzymes reflect their critical role in maintaining the integrity of the genetic material and ensuring proper cellular function.

RNR catalyzes the conversion of ribonucleotides to deoxyribonucleotides, which are the building blocks of DNA. RNR plays a crucial role in DNA replication and repair by providing the necessary precursors for DNA synthesis. The enzyme accomplishes this by reducing the 2'-hydroxyl group of ribose to a hydrogen atom, resulting in the conversion of ribonucleotides (i.e., ATP, GTP, CTP, and UTP) to their corresponding deoxyribonucleotides (i.e., dATP, dGTP, dCTP, and dTTP).



RNR is essential for the proliferation of all cells, including cancer cells, and is therefore a target for cancer chemotherapy. RNR is also a key enzyme in the synthesis of deoxyribonucleotides in bacteria and viruses, making it a potential target for the development of new antibiotics and antiviral drugs.

A.Hofer (2012): The first RNR activity was observed in the year 1950 by Swedish researcher Peter Reichard and coworkers, where they observed the conversion of ribonucleotides to deoxyribonucleotides.  Seven decades after its discovery, RNR is still a popular field to study in the scientific community. Perhaps, it is no exaggeration to say that RNR is the most interesting enzyme to study. Although it has been almost seven decades since the isolation of nucleotide reductase, RNR continues to surprise after all these years. The allosteric activity site (a-site), which functions as an on-off switch for the enzyme’s overall activity by binding ATP (activator) or dATP (inhibitor). The dNTP concentrations are probably optimized to minimize the mutation rate depending on the affinity of the DNA polymerase for different nucleotides. 19


Regulation of class I RNR. 
Class I RNRs are activated by binding ATP or inactivated by binding dATP to the activity site located on the RNR1 subunit. When the enzyme is activated, substrates are reduced if the corresponding effectors bind to the allosteric substrate specificity site. A = when dATP or ATP is bound at the allosteric site, the enzyme accepts UDP and CDP into the catalytic site; B = when dGTP is bound, ADP enters the catalytic site; C = when dTTP is bound, GDP enters the catalytic site. The substrates (ribonucleotides UDP, CDP, ADP, and GDP) are converted to dNTPs by a mechanism involving the generation of a free radical.25

Ribonucleotide reductase (RNR) is the only source for de novo production of the four deoxyribonucleoside triphosphate (dNTP) building blocks needed for DNA synthesis and repair. It is crucial that these deoxynucleotide triphosphates (dNTP) pools are carefully balanced since mutation rates increase when dNTP levels are either unbalanced or elevated. RNR is the major player in this homeostasis, and with its four different substrates, four different allosteric effectors  ( Wikipedia: allosteric regulation or control is the regulation of an enzyme by binding an effector molecule at a site other than the enzyme's active site) and two different effector binding sites, it has one of the most sophisticated allosteric regulations known today.24 

This regulation is achieved through multiple mechanisms, including allosteric regulation, protein-protein interactions, and post-translational modifications. For example, in E. coli, RNR activity is regulated by two small protein subunits, called SmlA and SmlB, which bind to the enzyme and inhibit its activity when dNTP levels are high. Additionally, RNR activity can be modulated by the presence of specific metabolites, such as ATP, which bind to allosteric sites on the enzyme and enhance or inhibit its activity.

The regulatory subunits of RNR enzymes do not directly recognize the level of DNA in the cell. Instead, they sense the levels of deoxyribonucleotide triphosphates (dNTPs). The mechanism by which the RNR enzymes sense the level of dNTPs is not fully understood, but several models have been proposed. One model suggests that the allosteric regulation of RNR enzymes by dNTPs involves direct binding of the dNTP to the RNR enzyme, leading to a conformational change that affects the activity of the enzyme. Another model suggests that the binding of dNTPs to the RNR enzyme changes the redox potential of the active site, altering the activity of the enzyme. The exact mechanism may vary among different organisms and RNR isoforms.

For the RNR enzyme to regulate the homeostasis of dNTPs in the cell, there must be a way for information about the cellular dNTP levels to be transmitted to the enzyme and processed by its regulatory mechanisms. This transmission of information can occur through a variety of mechanisms, including direct binding of dNTPs to the RNR enzyme or to its allosteric regulators, as well as signaling pathways that involve other proteins or molecules. Once the information is transmitted to the RNR enzyme, it can be processed through a series of regulatory mechanisms, such as post-translational modifications or protein-protein interactions, to modulate the activity of the enzyme and maintain the appropriate balance of dNTPs in the cell.

The regulatory mechanisms and information transmission systems involved in the regulation of RNR activity are interdependent and must be fully functional in order to perform their tasks effectively.

For example, the allosteric regulation of RNR activity by dNTPs requires the presence of specific binding sites on the enzyme and the ability of these sites to bind to dNTPs with high affinity. Similarly, the post-translational modifications and protein-protein interactions that regulate RNR activity rely on the presence of specific enzymes and proteins that can carry out these modifications and interactions.

If any of these regulatory mechanisms or information transmission systems are impaired or disrupted, it can lead to imbalances in the cellular dNTP pool, which can have serious consequences for DNA replication and repair, and ultimately for the survival and proliferation of the organism.

Therefore, it is important for the regulatory mechanisms and information transmission systems involved in the regulation of RNR activity to be fully functional and properly coordinated in order to maintain the appropriate balance of dNTPs in the cell and ensure proper DNA synthesis and repair.

Chabes and Thelander (2000): In order to maintain a balanced dNTP pool, RNR activity must be tightly regulated by multiple mechanisms, including transcriptional, post-transcriptional, translational, and post-translational control. These regulatory mechanisms are interdependent and must be fully functional in order to maintain the appropriate balance of dNTPs for DNA synthesis and repair. The allosteric regulation of RNR activity by dNTPs involves direct binding of the dNTP to the RNR enzyme or to its allosteric regulators, leading to a conformational change that affects the activity of the enzyme. The binding of dNTPs to the RNR enzyme can also alter the redox potential of the active site, modulating the activity of the enzyme. In addition, post-translational modifications and protein-protein interactions can regulate RNR activity by affecting enzyme localization, stability, and interactions with other proteins in the dNTP synthesis pathway. 27

Genes encoding RNR enzymes

The genes encoding the simplest RNR enzymes are typically organized in operons that contain other genes involved in DNA replication, repair, and recombination. The simplest RNR enzymes are classified into three classes based on the cofactor they use: class I enzymes use a glycyl radical, class II enzymes use a cobalamin (B12) cofactor, and class III enzymes use an iron-sulfur (FeS) cluster.

The gene that encodes for the large subunit of the RNR enzyme is typically named nrdA, while the gene that encodes for the small subunit is named nrdB or nrdD, depending on the organism. In some cases, the genes for both subunits are fused into a single gene, which is named nrdAB or nrdA/B.

The genes encoding class I enzymes are typically designated as nrdD, and they are usually located adjacent to nrdG and nrdE, which encode proteins involved in the activation of the glycyl radical cofactor. The nrdD gene encodes the catalytic subunit of the enzyme, which contains the active site that catalyzes the conversion of ribonucleotides to deoxyribonucleotides.

The genes encoding class II enzymes are typically designated as nrdJ, and they are usually located adjacent to nrdI and nrdH, which encode proteins involved in the activation of the cobalamin cofactor. The nrdJ gene encodes the catalytic subunit of the enzyme, which contains the active site that catalyzes the conversion of ribonucleotides to deoxyribonucleotides.

The genes encoding class III enzymes are typically designated as nrdG, and they are usually located adjacent to nrdD, which encodes the catalytic subunit of the enzyme. The nrdG gene encodes a small subunit that contains an FeS cluster and is involved in the generation of the tyrosyl radical that is required for catalysis by the enzyme.

The smallest genes expressing RNR enzymes are found in some bacteriophages (viruses that infect bacteria). These viruses have very compact genomes, and their RNR genes can be as small as 300-400 base pairs (bp), which is much smaller than the typical bacterial RNR genes that range from 2,000 to 4,000 bp. For example, the T4 bacteriophage, which infects Escherichia coli, encodes a small RNR enzyme consisting of only one polypeptide chain that is 153 amino acids long and has a molecular weight of 17.6 kDa. The gene encoding this enzyme is only 438 bp long and is designated nrdX. Despite their small size, these viral RNR enzymes are still functional and are important for the replication of the viral genome. Their compact size is thought to be an adaptation to the limited genome size of the virus, allowing them to maximize the amount of genetic information that can be stored in their genome while still retaining the essential function of RNR activity.

Processing of the RNR mRNA transcript once it is  transcribed

Once the mRNA transcript is transcribed, it undergoes several processing steps before it is translated into the RNR enzyme.

1. The first step is capping, where a modified guanine nucleotide is added to the 5' end of the mRNA. This protects the mRNA from degradation and helps to recruit the ribosome to the mRNA.
2. Next, the mRNA undergoes splicing in some organisms. In eukaryotes, some bacteria and archaea, the mRNA can contain introns - non-coding sequences that must be removed in order to generate a functional mRNA. The process of splicing involves the removal of introns and the ligation of exons, which are the coding regions of the mRNA.
3. The mature mRNA is then polyadenylated, where a string of adenosine nucleotides is added to the 3' end of the mRNA. Polyadenylation is a process that adds a sequence of adenine nucleotides (A's) to the 3' end of an mRNA molecule. This process is performed by an enzyme called poly(A) polymerase. The resulting modified mRNA molecule is called polyadenylated mRNA. The poly(A) tail can vary in length, but in most eukaryotic cells, it ranges from 100 to 200 nucleotides long. The poly(A) tail plays a critical role in the regulation of mRNA stability, transport, and translation. The addition of the poly(A) tail occurs after the RNA transcript is cleaved at a specific site downstream of the coding region. The poly(A) tail protects the mRNA from degradation by exonucleases, enzymes that break down RNA molecules from the ends. Additionally, the poly(A) tail is thought to facilitate the export of the mRNA from the nucleus, aid in the initiation of translation, and increase the efficiency of translation by promoting ribosome binding. Polyadenylation is a common feature of eukaryotic mRNA processing, but it also occurs in some bacterial and viral mRNA molecules.

Polyadenylation is a common feature of eukaryotic mRNA processing, but it does not occur in all cases. In some prokaryotic organisms, such as bacteria, mRNA molecules do not typically undergo polyadenylation. Instead, the 3' end of the mRNA is often processed by a ribonuclease enzyme that cleaves the RNA molecule after a specific sequence, which can vary depending on the organism. However, there are some exceptions where polyadenylation has been observed in bacterial mRNA. For example, in some species of bacteria, including Escherichia coli, polyadenylation can occur in certain mRNA molecules, such as those involved in stress response or regulatory functions. In general, polyadenylation is a more common feature of eukaryotic mRNA processing, but it is not universal and can vary depending on the organism and the specific mRNA molecule being processed.

Some studies have investigated the RNA metabolism of certain extremophilic organisms, such as thermophiles, which live in very high-temperature environments. These organisms have adapted to the extreme conditions of their environment by developing unique mechanisms for RNA processing and stability, which may differ from those of more typical organisms. In some cases, these mechanisms do not involve polyadenylation or other similar processing steps. Additionally, some studies have investigated the RNA metabolism of viruses, which are not technically considered to be alive but can still replicate and interact with host organisms. Many viruses have unique mechanisms for RNA processing and stability that do not involve polyadenylation or other similar processing steps. While polyadenylation is a common feature of RNA processing in most modern organisms, there is evidence to suggest that life can exist without this processing under certain conditions. However, it is important to note that the precise mechanisms of RNA processing and stability can vary widely between different types of organisms, and our understanding of these mechanisms is still evolving.

Some type of processing is necessary for RNA molecules to function properly, even if it is not polyadenylation or a similar mechanism. RNA molecules are typically synthesized as longer, precursor molecules that must undergo various processing steps to become functional. For example, in many organisms, precursor RNA molecules must be modified by removing certain sequences (such as introns) and adding certain chemical modifications (such as methyl groups or cap structures) to produce the final, mature RNA molecule. These processing steps are important for several reasons. First, they can help to ensure that the RNA molecule is functional and can perform its intended role in the cell. Second, they can help to regulate gene expression by influencing the stability, localization, or translation efficiency of the RNA molecule. Finally, they can provide a mechanism for cells to respond to environmental or developmental cues by altering the processing or stability of specific RNA molecules. While the precise processing steps required for RNA molecules can vary widely between different types of organisms and RNA molecules, it is generally true that some type of processing is necessary for RNA molecules to function properly in cells.

The origin of mRNA processing is not as well-understood as the origin of life itself, as the molecular mechanisms involved in RNA processing are complex and still the subject of ongoing research. However, there are several proposals and hypotheses related to the origin of mRNA processing in early life. One hypothesis is that early RNA molecules were not processed in the same way that modern mRNA is processed. Instead, it is possible that early RNA molecules were shorter and more primitive, lacking some of the more complex structures and modifications found in modern mRNA. Over time, these processes evolved and became more complex, RNA processing mechanisms would have become more sophisticated to enable greater control over gene expression and protein synthesis. There is no direct empirical evidence to support the idea that simpler forms of RNA processing would have worked in early life.
Studies of the RNA molecules found in modern organisms have shown that many of the complex structures and modifications found in modern mRNA are not strictly necessary for RNA to function. In some cases, RNA molecules with simpler structures and fewer modifications can still carry out their biological functions.  The evolution of complex enzymes capable of processing RNA would have been a significant step in the development of modern gene expression mechanisms. The emergence of such enzymes would have required the selection of specific genetic sequences capable of coding for the necessary protein structures, as well as the optimization of complex biochemical pathways for RNA processing. An only feasible explanation would be that the development of RNA processing mechanisms would have been a gradual, stepwise process that occurred over a long period of time. Some researchers have proposed that the initial steps in this process may have involved the emergence of simple RNA-binding proteins that could help to stabilize RNA molecules and protect them from degradation. Over time, these proteins could have become more complex and developed the ability to modify RNA molecules in specific ways, leading to the evolution of the more complex RNA processing enzymes seen in modern organisms.  The evolution of RNA processing mechanisms would have been a key step in the development of modern gene expression and protein synthesis, and this process would have had to involve a combination of genetic and biochemical changes occurring over a long period of time.

The simplest known RNA-binding proteins are typically composed of one or a few RNA-binding domains (RBDs), which are protein domains that specifically recognize and bind to RNA molecules. These RBDs can be found in a wide range of proteins, from simple RNA chaperones to more complex RNA-binding proteins involved in various aspects of gene expression and regulation. One example of a simple RNA-binding protein is the S1 ribosomal protein, which is found in many prokaryotic and eukaryotic ribosomes and plays a role in stabilizing the mRNA molecule during translation. The S1 protein has a single RNA-binding domain that recognizes specific RNA sequences in the 5' untranslated region (UTR) of mRNA. Another example is the Hfq protein, which is found in many bacteria and is involved in the regulation of mRNA stability and translation. Hfq has a single RNA-binding domain and functions as a chaperone to facilitate interactions between small regulatory RNAs and their mRNA targets. While these proteins are relatively simple compared to the more complex RNA-binding proteins found in modern organisms, they provide important insights into the evolution of RNA-protein interactions and the development of more complex RNA processing mechanisms over time.

The size of the smallest known RNA-binding domain (RBD) is around 50 amino acids, which corresponds to a molecular weight of approximately 5-6 kDa. An example of such a small RBD is the RNP-1 motif, which is found in a variety of RNA-binding proteins and is characterized by a conserved sequence motif that forms a beta-alpha-beta fold structure that is capable of binding to RNA. Other small RNA-binding domains include the K-homology (KH) domain, which is typically around 70 amino acids in size, and the RNA recognition motif (RRM), which is around 90 amino acids in size. These small RBDs are often found in larger RNA-binding proteins that contain multiple RBDs and perform more complex functions. It's generally thought that around 40-50 amino acids is the minimum length required for a functional RBD, as this length is sufficient to form the basic structural elements required for RNA binding, such as alpha helices and beta sheets. If the RBD were much smaller than this, it's possible that it would not be able to form the necessary structures and would therefore be non-functional.

The probability of a random sequence of amino acids spontaneously forming a functional protein is very low. The vast majority of possible amino acid sequences do not fold into stable, functional proteins, so the odds of randomly generating a functional protein are typically estimated to be around 1 in 10^77 or lower. Some Origin of Life ( OoL) researchers claim that the emergence of functional proteins in early life could have been facilitated by a number of factors, including non-random chemical selection of amino acids and the availability of prebiotic conditions that favored the formation and stabilization of functional protein structures. So while the odds of a small protein becoming functional by chance alone are low, it's possible that early life on Earth was able to overcome these odds through a combination of chemical and environmental factors. Non-random chemical selection of amino acids however would imply a directed process, which is not consistent with the concept of natural selection.

A typical cop-out in face of these huge problems is to say: " Progress has been made in recent years in areas such as prebiotic chemistry, the role of RNA in early life, and the potential for life to arise in other environments beyond Earth. While there is still much to learn, the scientific community continues to make strides in understanding the origins of life."

4. Once the mRNA has undergone these processing steps, it is exported from the nucleus (if present) and can be translated into the RNR enzyme. The mRNA is recognized by ribosomes, which read the nucleotide sequence and translate it into the corresponding amino acid sequence, ultimately producing the functional RNR enzyme.

Posttranslational modifications after translation

Once the RNR protein subunits are synthesized, they may undergo additional post-translational modifications to become fully functional. The large subunit (NrdA) may be subject to several modifications, including the formation of disulfide bonds, the addition of a metal cofactor, and proteolytic cleavage to form an active enzyme complex.

In some organisms, the NrdA subunit may also be subject to feedback inhibition, where the product of the reaction (deoxyribonucleotides) can bind to the active site and inhibit further enzyme activity. This feedback inhibition may be regulated by additional post-translational modifications to the protein, such as phosphorylation or allosteric regulation.

The small subunit (NrdB or NrdD) may also undergo additional modifications, such as the binding of a metal cofactor or the formation of disulfide bonds, to become fully functional. In some cases, the small subunit may also be subject to proteolytic cleavage or other modifications to regulate enzyme activity.

Overall, post-translational modifications play a critical role in regulating the activity and function of the RNR enzyme, and allow for tight control over the synthesis of deoxyribonucleotides needed for DNA replication and repair.

Formation of disulfide bonds in RNR strands

The formation of disulfide bonds is necessary for the proper folding and stability of many proteins, including RNR enzymes. Disulfide bonds are covalent bonds that form between two cysteine residues in a protein, and they can help to stabilize the protein structure by providing a bridge between different parts of the protein. In RNR enzymes, the disulfide bonds help to maintain the correct conformation of the enzyme, which is essential for its activity. Without these bonds, the enzyme may not function properly or may be degraded more quickly.  The number of disulfide bonds required for the RNR enzyme structure can vary depending on the organism and the specific subunit of the enzyme. For example, in E. coli, the RNR enzyme contains four subunits: alpha 2, beta 2, gamma, and delta. The alpha 2 and beta 2 subunits each contain two disulfide bonds, while the gamma and delta subunits do not contain any disulfide bonds. Therefore, in this case, a total of four disulfide bonds are necessary for the full RNR enzyme structure. However, in other organisms, the number and placement of disulfide bonds may differ. The formation of disulfide bonds is catalyzed by a group of enzymes called oxidoreductases, which are also known as disulfide isomerases. These enzymes catalyze the transfer of electrons between cysteine residues, leading to the formation of disulfide bonds. The process involves the oxidation of two cysteine thiol groups to form a disulfide bond (S-S bond) and the reduction of a disulfide bond to two cysteine thiol groups. In the case of RNR enzymes, the formation of disulfide bonds is important for the correct folding and stability of the protein, as well as for the regulation of its activity. The specific mechanism by which disulfide bonds are formed in RNR enzymes may vary depending on the organism and the specific RNR enzyme in question. Enzymes that catalyze disulfide bond formation typically recognize specific amino acid sequences or structural features in the protein that are involved in forming the disulfide bond. These enzymes, called protein disulfide isomerases (PDIs), contain specific domains or motifs that recognize and bind to these sequences or structural features. In the case of RNR enzymes, the formation of the disulfide bond is thought to be guided by the presence of specific cysteine residues in the protein sequence. These cysteine residues are typically located in regions of the protein that are important for stabilizing the overall structure of the enzyme or for forming critical active sites where substrate binding and catalysis occur. The PDIs recognize these cysteine residues and catalyze the formation of disulfide bonds between them. The exact mechanism by which PDIs recognize specific cysteine residues and catalyze disulfide bond formation is still an area of active research. Disulfide bonds are important for stabilizing the structure of the RNR enzymes, and without them, the enzymes could be more susceptible to degradation and may not function properly.

The process of disulfide bond formation is closely monitored and regulated by the cell. Cells have a number of enzymes that are responsible for catalyzing disulfide bond formation and rearrangement, as well as for monitoring the quality of the disulfide bonds that are formed. One of the key enzymes involved in disulfide bond formation is protein disulfide isomerase (PDI). PDI is a chaperone protein that helps to fold and stabilize newly synthesized proteins, including RNR enzymes. It also catalyzes the formation and rearrangement of disulfide bonds. In addition, cells have a quality control mechanism that monitors the folding and stability of newly synthesized proteins. Proteins that are misfolded or have improperly formed disulfide bonds are recognized by chaperone proteins and targeted for degradation by the cell's protein degradation machinery. This helps to ensure that only properly folded and functional proteins are present in the cell.

Metallocofactor assembly

In RNR enzymes, the addition of metal cofactors is typically performed through a process known as metallocofactor assembly. The specific details of this process can vary depending on the organism and the type of metal involved, but in general, it involves the coordination of the metal ion with specific amino acid residues in the protein.

For example, in class I RNR enzymes, which use a diferric-tyrosyl radical (Fe^III-Fe^III-Y•) cofactor, the metallocofactor assembly involves the binding of two iron ions to a specific site on the protein, followed by the binding of a tyrosine residue to one of the iron ions. This tyrosine residue is then oxidized to form a tyrosyl radical, which is stabilized by the adjacent iron ions.

The probability of a non-evolutionary, non-intelligent mechanism finding the right spot for binding of metal ions to a specific site on a protein is extremely low. The binding of metal ions to a specific site on a protein involves specific chemical interactions, which require precise spatial and electrostatic arrangements.  The active site contains specific amino acid residues that are able to coordinate the metal ions and hold them in place. These residues often contain functional groups like cysteine or histidine, which have the ability to bind metal ions through the donation of electrons. The spatial arrangement of these residues is critical to ensure that the metal ions are held in the correct orientation and with the proper distance between them to enable their catalytic activity. In addition to the specific amino acid residues involved in metal coordination, other surrounding residues can also play a role in electrostatic interactions that help to stabilize the active site and enhance its catalytic activity. These residues may interact with the metal ions or other charged or polar groups involved in the reaction through hydrogen bonding, van der Waals interactions, or other mechanisms. The odds of finding the right spot for metal ion binding and encoding the information to find it through random processes alone are extremely low, given the vastness of the sequence space and the complexity of the required interactions. The probability of such an event happening by chance is considered to be exceedingly low, and would require a vast amount of time and a vast number of trials, far beyond what is thought to be possible in the age of the universe. Therefore, many scientists argue that the origin of such complex biological structures requires non-random mechanisms, such as intelligent design or directed evolution.

In class II RNR enzymes, which use a cobalt or manganese ion as the metal cofactor, the metallocofactor assembly process is somewhat different. In these enzymes, the metal ion is typically bound to a specific site on the protein, followed by the coordination of additional ligands to the metal ion to stabilize it in its active form.

There would be many more non-functional configurations possible, than functional ones, especially considering the complexity of the coordination required for the metallocofactor assembly process in RNR enzymes. The chance of finding the correct configuration by chance alone is exceedingly low, and it is unlikely that this process could have arisen through unguided natural processes alone. This is one reason why some scientists argue that there must be some form of intelligent design or intervention in the origin of life and the development of complex biological systems.

Interdependence in biological systems is a hallmark of design

DNA is the genetic material, that is used in all known life forms. No exception.  The origin of DNA is an Origin of Life problem.  Ribonucleotide reductase (RNR) is the only source for de novo production of the four deoxyribonucleoside triphosphate (dNTP) building blocks needed for DNA synthesis and repair. RNR catalyzes the conversion of ribonucleotides to deoxyribonucleotides, which are the building blocks of DNA. The origin of RNR enzymes is therefore as well an Origin of Life problem. They are among the most sophisticated and complex enzymes known.  Not only are they involved in de-novo production, but also monitor, control, and regulate the appropriate level of DNA in the cell, which is essential for cell survival.  To perform their action,  they sense the levels of deoxyribonucleotide triphosphates (dNTPs) in the cell. There is a way for information about the cellular dNTP levels to be transmitted to the enzyme and processed by its regulatory mechanisms. This transmission of information can occur through a variety of mechanisms, including direct binding of dNTPs to the RNR enzyme or to its allosteric regulators, as well as signaling pathways that involve other proteins or molecules. Once the information is transmitted to the RNR enzyme, it can be processed through a series of regulatory mechanisms, such as post-translational modifications or protein-protein interactions, to modulate the activity of the enzyme and maintain the appropriate balance of dNTPs in the cell. Chabes and Thelander (2000): 

These regulatory mechanisms are interdependent and must be fully functional to maintain the appropriate balance of dNTPs for DNA synthesis and repair.   Interdependence in a biological system means that the components of the system are interdependent and rely on each other to function properly. If one or more components are impaired or disrupted, it can lead to a breakdown of the system and an inability to perform its function effectively. In biological systems, it is often the case that the interdependence of components leads to a "all-or-nothing" or "threshold" effect, where the system requires all components to be functioning properly to perform its function effectively. The individual units or components of a biological system only have a function when integrated into the larger system. Biological systems are composed of multiple components that work together in a coordinated manner to achieve a specific function. For example, in the case of RNR regulation, the individual subunits of the enzyme only have a function when integrated into the larger enzyme complex, which requires multiple subunits to function properly. Similarly, the regulatory mechanisms and information transmission systems that control RNR activity only have a function when integrated into the larger system of dNTP synthesis and DNA replication and repair. Therefore, the function of individual components in biological systems is often dependent on their integration into larger systems, where they work together in a coordinated manner to achieve a specific biological function.  

Interdependent systems are a fundamental feature of biological organisms, and many different systems in the cell are interdependent and essential for life. For example, in addition to the regulation of RNR activity and the maintenance of the dNTP pool, many other interdependent systems in the cell are essential for life, such as the regulation of gene expression, protein synthesis and degradation, energy metabolism, and cell signaling. Each of these systems relies on the interdependence of multiple components and regulatory mechanisms to function properly, and disruption of any one of these systems can have serious consequences for the cell and the organism as a whole. Therefore, the interdependence of biological systems is a key feature of life, reflecting the complex and highly integrated nature of living organisms.

The emergence of the first living organisms would have required the formation of complex biochemical systems, including the RNR enzymes, which are essential for DNA synthesis and repair.

A naturalistic hypothesis is that the first living organisms emerged through a process of chemical evolution, where simple organic molecules combined and interacted to form more complex molecules and eventually self-replicating systems. This process would have taken place in prebiotic environments, such as hydrothermal vents or in the early Earth's atmosphere. That would have required the formation of complex biochemical systems, including the RNR enzymes. There are no plausible models of the specific naturalistic mechanisms that could have led to the emergence of these systems. The direct observation of the emergence of complex interdependent systems is challenging, as it would require tracking the development of these systems over long periods of time, which is not feasible. A hypothesis presented is that individual parts were co-opted in the environment.  RNR enzymes would have involved the co-option of pre-existing biochemical pathways or enzymes that were adapted for a new function in DNA synthesis and repair.  This hypothesis, however, faces considerable hurdles.

For a working biological system to be built, the five following conditions would all have to be met:
C1: Availability. Among the parts available for recruitment to form the system, there would need to be ones capable of performing the highly specialized tasks of individual parts, even though all of these items serve some other function or no function.
C2: Synchronization. The availability of these parts would have to be synchronized so that at some point, either individually or in combination, they are all available at the same time.
C3: Localization. The selected parts must all be made available at the same ‘construction site,’ perhaps not simultaneously but certainly at the time, they are needed.
C4: Coordination. The parts must be coordinated in just the right way: even if all of the parts of a system are available at the right time, it is clear that the majority of ways of assembling them will be non-functional or irrelevant.
C5: Interface compatibility. The parts must be mutually compatible, that is, ‘well-matched’ and capable of properly ‘interacting’: even if subsystems or parts are put together in the right order, they also need to interface correctly.
( Agents Under Fire: Materialism and the Rationality of Science, pgs. 104-105 (Rowman & Littlefield, 2004). HT: ENV.)

Those conditions make the co-option of parts to form a working biological system quite challenging, especially considering the vast number of specialized parts that would need to be coordinated and integrated. The probability of all the necessary parts being available, synchronized, localized, coordinated, and compatible at the same time and place is extremely low, making the origin of complex biological systems through gradual co-option of parts highly unlikely.  On the other hand, intelligent agents are capable of designing and building complex systems that exhibit the characteristics mentioned in the five conditions. Intelligent agents can identify the necessary components, synchronize their availability, localize them, coordinate them, and ensure their interface compatibility, to create complex systems that function as intended. There is ample evidence in fields such as engineering, computer science, and architecture that intelligence is capable of designing and building complex systems that require coordination and interdependence. We only have direct evidence of intelligence being capable of instantiating complex interdependent systems, while naturalistic explanations for the origin of such systems remain hypothetical and unproven.


Ribonucleotide Reductase Converts Ribonucleotides to Deoxyribonucleotides. D. Voet et.al. (2016): Deoxyribonucleotides are synthesized from their corresponding ribonucleotides by the reduction of their C2′ position rather than by their de novo synthesis from deoxyribose-containing precursors.


Enzymes that catalyze the formation of deoxyribonucleotides by the reduction of the corresponding ribonucleotides are named ribonucleotide reductases (RNRs). RNRs are one of the most essential enzymes of life. There are three classes of RNRs, which differ in their prosthetic groups. ( Wikipedia: A prosthetic group is the non-amino acid component that is part of the structure of the heteroproteins or conjugated proteins, being tightly linked to the apoprotein.) They all replace the 2′-OH group of ribose with H via a free-radical mechanism involving a thiyl radical. 

A. A. Burnim et.al.,(2022): Ribonucleotide reductases (RNRs) are used by all free-living organisms and many viruses to catalyze this essential step in the de novo biosynthesis of DNA precursors. 8

What-when-how: Different organisms employ widely different forms of three different classes of the enzyme. All classes of RNR share two exceptional features:

1. The polypeptide chain of the active enzyme harbors a free radical amino acid residue that participates in the catalytic process; the mechanism for radical generation sets the classes apart
2. The specificity toward the four ribonucleotides is tightly controlled by allosteric effects that are remarkably similar for the three classes.7

Daniel Lundin (2015): It is remarkable that RNR uses some of the most potent metals in redox chemistry. All RNRs use radical chemistry to catalyze this challenging reaction.9
A.Hofer (2011): It is crucial that these dNTP pools are carefully balanced since mutation rates increase when dNTP levels are either unbalanced or elevated. RNR is the major player in this homeostasis, and with its four different substrates, four different allosteric effectors, and two different effector binding sites, it has one of the most sophisticated allosteric regulations known today. Allosteric regulation of RNRs affects both substrate specificity and overall activity. The s-site binds dNTPs and determines which nucleotide will be reduced at the active site to ensure balanced levels of the four deoxyribonucleotides dNTPs in the cell.10
Soo-Cheul Yoo (2009): Ribonucleotide reduction is the only pathway for de novo synthesis of deoxyribonucleotides in extant organisms. This chemically demanding reaction, which proceeds via a carbon-centered free radical. The mechanism has been deemed unlikely to be catalyzed by a ribozyme, creating an enigma regarding how the building blocks for DNA were synthesized at the transition from RNA to DNA-encoded genomes. 11

Comment: Here we have a classic chicken and egg problem.  RNR enzymes are required to make DNA. DNA is however required to make RNR enzymes. What came first ??  We can conclude with high certainty that this enzyme buries any RNA world hypothesis and any possibility of transition from  RNA to DNA world scenarios.

There are 3 classes of RNR enzymes

There are three classes of RNR enzymes, which differ in their structure and mechanism of action.

Class I RNR enzymes are found in eukaryotes, bacteria, and viruses. They use a radical mechanism to generate a stable tyrosyl radical on the enzyme, which then abstracts an electron from the substrate to initiate the reduction reaction. Class I RNR enzymes require a protein called R1 to provide the catalytic site for the reduction of ribonucleotides. R1 contains a dinuclear metal center composed of iron and tyrosine residues that are essential for the activity of the enzyme.

Class II RNR enzymes are found in bacteria and archaea. They use a different radical mechanism to generate a stable glycyl radical on the enzyme, which then abstracts an electron from the substrate. Unlike Class I RNR enzymes, Class II RNR enzymes do not require a separate protein for their activity, and the active site is located entirely within the enzyme.

Class III RNR enzymes are only found in aerobic bacteria and archaea. They use a radical mechanism similar to Class I RNR enzymes, but the reaction is initiated by a flavodoxin protein instead of a tyrosyl radical. Class III RNR enzymes are not well understood, and their function in these organisms is not yet clear.

E. Torrents (2014): Currently, three different RNR classes have been described (I, II, and III), and class I is further subdivided into Ia, Ib, and Ic. All three RNR classes share a common three-dimensional protein structure at the catalytic subunit and a highly conserved α/β barrel structure in the active site of the enzyme. In addition, the two potential allosteric centers (specificity and activity) are highly conserved among the different RNR classes, although in class Ib, and some class II RNRs activity allosteric site is absent.20 

Daniel Lundin (2009):The significant differences between RNRs exist notably in cofactor requirements, subunit composition and allosteric regulation. These differences result in distinct operational constraints (anaerobicity, iron/oxygen dependence, and cobalamin dependence), and form the basis for the classification of RNRs into three classes. 22

T. B. Ruskoski (2021): More recently, remarkably diverse bioinorganic and radical cofactors have been discovered in class I RNRs from pathogenic microbes. These enzymes use alternative transition metal ions, such as manganese, or posttranslationally installed tyrosyl radicals for initiation of ribonucleotide reduction.21

Why are there 3 classes? 

While all three classes of RNR enzymes catalyze the same fundamental reaction of converting ribonucleotides to deoxyribonucleotides, they perform this reaction in different ways and under different conditions. This diversity is likely due to the varying environments and metabolic needs of the organisms in which these enzymes are found.

For example, Class I RNR enzymes are found in a wide range of organisms, including eukaryotes, bacteria, and viruses, and are essential for DNA replication and repair. Class II RNR enzymes, on the other hand, are only found in bacteria and archaea, and they are generally more resistant to oxidative stress than Class I enzymes, which may be important for their survival in harsh environments. Class III RNR enzymes are found only in aerobic bacteria and archaea and are thought to play a role in the regulation of iron homeostasis.

In addition, the different classes of RNR enzymes have distinct structural and mechanistic features that may make them more or less suitable for different types of cellular processes. For example, Class I RNR enzymes require a separate protein for their activity, which may provide an additional level of regulation or allow for more precise control of the enzyme's activity. In contrast, Class II RNR enzymes are self-contained and may be better suited for rapid responses to changing environmental conditions.

Class II RNR enzymes found in bacteria and archaea function in environments with high levels of oxidative stress. Oxidative stress refers to an imbalance between the production of reactive oxygen species (ROS) and the cell's ability to detoxify them. ROS are highly reactive molecules that can damage cellular components such as proteins, lipids, and DNA, leading to cellular dysfunction and death. ROS are produced as byproducts of various metabolic processes, including respiration and photosynthesis, and their levels can increase in response to environmental stressors such as exposure to UV radiation or toxins.

Class II RNR enzymes found in bacteria and archaea are adapted to function in environments with high levels of oxidative stress. These enzymes have unique structural and mechanistic features that allow them to withstand and repair the damage caused by ROS. For example, they have a unique mechanism for generating a glycyl radical, which is used to initiate the reduction reaction and is highly resistant to oxidation. Additionally, Class II RNR enzymes have been shown to interact with antioxidant enzymes, such as thioredoxins and glutaredoxins, which can help to reduce ROS levels and prevent oxidative damage.

The ability of Class II RNR enzymes to function in environments with high levels of oxidative stress is essential for the survival and adaptation of bacteria and archaea in harsh environments. Reducing ROS is an additional function of Class II RNR enzymes, in addition to their primary function of transforming RNA to DNA. They can be considered multifunctional enzymes, as they have more than one function in the cell.  Some studies have suggested that Class II RNR enzymes may have a role in the regulation of gene expression, particularly in response to stress or nutrient availability. Other studies have suggested that Class II RNR enzymes may have a role in the production of secondary metabolites or in the metabolism of xenobiotics (foreign compounds). However, these proposed functions are still the subject of ongoing research and are not yet fully understood.

What are the environments that require organisms with Class II RNR enzymes?

Class II RNR enzymes are required by organisms that live in harsh environments where oxygen is scarce or absent, such as deep-sea hydrothermal vents, anaerobic sediments, or inside the guts of some animals. These environments are characterized by low levels of oxygen, high levels of toxic compounds, extreme temperatures, and high pressures.

In these environments, Class II RNR enzymes are necessary for the synthesis of deoxyribonucleotides, the building blocks of DNA. These enzymes use a different mechanism than Class I RNR enzymes to generate the free radical needed to initiate the reaction, which does not require oxygen. This allows organisms to synthesize DNA even in the absence of oxygen.

Some examples of organisms that require Class II RNR enzymes include anaerobic bacteria such as Clostridium species, which live in the gut of animals and are involved in the breakdown of organic matter, and archaea such as Pyrococcus furiosus, which live in hot environments such as deep-sea hydrothermal vents and use Class II RNR enzymes to synthesize DNA in the absence of oxygen.

How are  Class II RNR enzymes distinct from the other two classes?

Class II RNR enzymes are distinct from the other two classes (Class I and Class III) in several ways:

Structure: Class II RNR enzymes have a completely different protein structure than Class I and III RNR enzymes. They consist of a single protein subunit, unlike Class I and III RNR enzymes, which are composed of multiple subunits.

Oxygen-independent: Class II RNR enzymes do not require oxygen to generate the free radical needed to initiate the reaction, unlike Class I RNR enzymes which use oxygen as a co-substrate, and Class III RNR enzymes which require a protein called AdoCbl (Adenosylcobalamin) to generate the free radical.

Metallocofactor: Class II RNR enzymes contain a different metallocofactor (metal ion-containing non-protein component) than the other two classes. Specifically, Class II RNR enzymes use a non-heme iron center with a tyrosyl radical, whereas Class I RNR enzymes use a di-iron center, and Class III RNR enzymes use AdoCbl.

Class II RNR enzymes are believed to have a different origin than the other two classes. Class II RNR enzymes appear to have a distinct origin and are only found in certain bacterial and archaeal species.

These enzymes are highly resistant to oxidation and have unique features that allow them to function in such conditions. In contrast, Class I RNR enzymes found in eukaryotes, bacteria, and viruses are optimized to function in environments with lower levels of oxidative stress and have different mechanisms to facilitate their activity. Similarly, Class III RNR enzymes are specialized to function in aerobic bacteria and archaea and have unique mechanisms to allow them to operate in these environments.

The differences in the environments in which these enzymes function can be attributed to a variety of factors, including the presence of different reactive oxygen species, variations in pH and temperature, and variations in the availability of cofactors and substrates. The mechanisms employed by each class of RNR enzymes have evolved to optimize their activity in their respective environments and to ensure that they can perform their essential functions under the appropriate conditions.

In summary, the unique features of the different classes of RNR enzymes allow them to function optimally in different environments, depending on the organism in which they are found. This specialization is necessary for the efficient and effective operation of cellular processes and highlights the importance of the environment in shaping the function and evolution of biological molecules.

Independent origin of the three RNR classes

Class II RNR enzymes most likely do not share a common ancestor with the other two classes of RNR enzymes, but emerged separately. This is supported by several lines of evidence, including their distinct protein structure, metallocofactor, and mechanism of action. 28 The history and origin of RNR enzymes are intimately connected to the broader question of the origin of life on Earth since these enzymes play a critical role in the synthesis and maintenance of genetic material in all living organisms, The three classes of Ribonucleotide Reductase (RNR) enzymes most likely have an independent and unique trajectory of origin. Class I RNR enzymes use a metallocofactor, a diferric-tyrosyl radical, to catalyze the conversion of nucleotides into deoxynucleotides. Class II RNR enzymes have a different structure and mechanism than the other two classes. They use a stable tyrosyl radical, rather than a diferric-tyrosyl radical, to initiate the nucleotide reduction reaction. Class III RNR enzymes were discovered relatively recently and have a unique mechanism that involves a stable glycyl radical and do not require any metals or cofactors for activity. The mechanisms of horizontal gene transfer, gene duplication, and convergent evolution are not adequate to explain all of the dissimilarities between the three classes of RNR enzymes.

The three metal RNR Co-factors

Class I of Ribonucleotide reductases occurs in aerobically thriving organisms including humans uses oxygen activated by a dinuclear iron center to convert a tyrosine residue into a radical.
Class II is the coenzyme B12-dependent reductase
Class III contains an extremely oxygen-sensitive glycyl radical, which is generated with the aid of S-adenosylmethionine (SAM).

All three types, however, use a thiyl radical at the active site and act by an almost identical mechanism.

All three types use a thiyl radical at the active site and act by an almost identical mechanism.

Lander, E. S (2001): Different classes of RNR's have intriguing sequence “motifs” involving cysteines that appear to be important for the catalysis (in Escherichia coli, Cys-439, the radical site, and Cys-225 and Cys-462, which delivers two electrons and a proton). These motifs offer tantalizing suggestions that all RNRs are related by common ancestry but underwent divergent evolution so massive that only traces of evidence for homology remain in the sequences themselves. These motifs are inadequate to provide a statistically significant case for homology, however, and motifs are notoriously inadequate for confirming homology in general. 29

Comment: To claim common ancestry, in this case, is an ad-hoc assertion. Truth said, science is unable to infer a reasonable scenario out of the evidence, and all it can do, is resort to made-up stories, which bear no credibility. The best and most straightforward explanation is that a creator made RNRs, and equipped each of them with different ways to perform the same function.

RNR structure

Class I RNR enzymes

Class I ribonucleotide reductase (RNR) enzymes are composed of two subunits, RRM1 and RRM2.

RRM1 subunit is a large protein consisting of approximately 800 amino acids. It contains two domains, the N-terminal domain and the C-terminal domain. The N-terminal domain is responsible for binding to the small subunit (RRM2) and contains a zinc finger motif that is involved in protein-protein interactions. The C-terminal domain contains the active site for ribonucleotide reduction and is composed of two subdomains: the substrate-binding subdomain and the radical-generating subdomain.

RRM2 subunit is a smaller protein consisting of approximately 350 amino acids. It contains a single domain with a unique structure known as the RNR-specificity loop. This loop is responsible for determining the specificity of the enzyme for the different ribonucleotides and contains a conserved tyrosine residue that plays a critical role in the catalytic mechanism of the enzyme.

The tertiary structure of the RNR enzyme is complex and involves the interaction of the two subunits. RRM1 and RRM2 form a heterodimeric enzyme complex that is regulated by the binding of different allosteric effectors. In the absence of allosteric effectors, RNR is in an inactive state, but upon binding of allosteric effectors, the enzyme complex undergoes conformational changes that allow for activation of the enzyme and subsequent ribonucleotide reduction.


Quaternary structure of the active holoenzyme complex in class I RNR (PDB accession code 6W4X). Insets show the location of the active site in the catalytic α subunit (middle top) and the metallo- or radical cofactor (middle bottom and far right) in the β subunit. 30

Overall, the structure of class I RNR enzymes is highly conserved across species and is essential for the de novo synthesis of deoxyribonucleotides, which are critical for DNA synthesis and cell division.

Class II RNR enzymes

The structure of class II RNR enzymes is unique and distinct from class I enzymes. These enzymes are composed of a large α subunit and a smaller β subunit, and the two subunits form a stable heterodimeric complex.

The α subunit of class II RNR enzymes contains two domains: the N-terminal domain and the C-terminal domain. The N-terminal domain contains the di-iron center, which is responsible for ribonucleotide reduction. The C-terminal domain is responsible for binding to the β subunit and contains a loop structure known as the specificity loop, which determines the specificity of the enzyme for different ribonucleotides.

The β subunit of class II RNR enzymes contains a single domain that is responsible for binding to the α subunit. It contains a conserved cysteine residue that is involved in the regulation of the enzyme.

The tertiary structure of class II RNR enzymes is highly conserved across species and is critical for enzyme function. The α and β subunits form a heterodimeric enzyme complex that is regulated by several allosteric effectors. Binding of allosteric effectors causes conformational changes in the enzyme complex that allow for activation of the enzyme and subsequent ribonucleotide reduction.

Overall, the structure of class II RNR enzymes is unique and distinct from class I enzymes, but both classes of enzymes are essential for the de novo synthesis of deoxyribonucleotides and DNA replication.

Class III RNR enzymes

Class III ribonucleotide reductase (RNR) enzymes are found in bacteriophages and some bacteria. These enzymes use a glycyl radical to reduce ribonucleotides, similar to class I RNR enzymes.

The structure of class III RNR enzymes is unique and distinct from class I and class II enzymes. These enzymes are composed of a single polypeptide chain that contains three domains: the N-terminal domain, the central domain, and the C-terminal domain.

The N-terminal domain of class III RNR enzymes contains a glycyl radical that is essential for enzyme function. The glycyl radical is generated by a radical SAM (S-adenosylmethionine) enzyme and is stabilized by the protein environment.

The central domain of class III RNR enzymes contains a conserved cysteine residue that is involved in the regulation of the enzyme.

The C-terminal domain of class III RNR enzymes contains a cluster of iron-sulfur (Fe-S) clusters that are involved in electron transfer and ribonucleotide reduction.

The tertiary structure of class III RNR enzymes is critical for enzyme function and involves the interaction of the three domains. The glycyl radical in the N-terminal domain is stabilized by the protein environment, and the central and C-terminal domains are involved in electron transfer and ribonucleotide reduction.

Overall, the structure of class III RNR enzymes is unique and distinct from class I and class II enzymes, but all three classes of enzymes are essential for the de novo synthesis of deoxyribonucleotides and DNA replication.

While all three classes of RNR enzymes share a common function, their proteic architecture is quite distinct from one another. 

RNR uses radical chemistry to catalyze the reduction of each NTP. How the enzyme generates this radical, the type of cofactor and metal required, the three-dimensional structure of this enzyme complex and the dependence of oxygen are all characteristics that are considered when classifying RNRs.

The X-ray structures of the R1 catalytic component and the R2 di-iron component of the class I RNR from E. coli were determined . The catalytic subunit was found to be a novel 10-stranded α/β-barrel with a loop that hosts the thiyl radical protruding into its center.  Despite a lack of sequence similarity with other ribonucleotide reductases, all ribonucleotide reductases would have similar catalytic subunits, reflecting their similar catalytic strategies. The structure of the catalytic subunit of the class III enzyme revealed the characteristic 10-stranded α/β-barrel with a central loop bearing the thiyl radical precursor. Many features of the structure reported are remarkably similar to the structures of the catalytic subunits of the class I and class III enzymes, even though there is <10% sequence homology among them. The similarities and contrasts with the enzymes of other classes have much to tell us about all ribonucleotide reductases.

Comment: The three classes of RNR enzymes have the same catalytic activity, but different amino acid sequences to reach the same result. Science has no good explanations for the divergence.

RNR Mechanism  and reaction

The mechanism of ribonucleotide reductase (RNR) can vary depending on the class of the enzyme.  Each of the three classes has a distinct mechanism for converting ribonucleotides to deoxyribonucleotides.

The mechanism in Class I RNR enzymes

Class I RNR enzymes use a free radical mechanism. The free radical in RNR enzymes is generated through a specific reaction that involves the reduction of a disulfide bond in the enzyme's active site by a cysteine residue. The reaction in which the free radical in RNR enzymes is generated is a multi-step process that can be divided into two stages: initiation and propagation.

Initiation:

The RNR enzyme contains a disulfide bond between two cysteine residues in the active site. The two cysteine residues in the active site of RNR enzymes are typically recruited from the enzyme's own polypeptide chain. During the synthesis of the enzyme, the amino acid sequence of the polypeptide chain includes these cysteine residues in a specific location within the enzyme's three-dimensional structure, which ultimately forms the enzyme's active site. In some cases, the cysteine residues may be supplied by a separate protein that interacts with the RNR enzyme, but this is less common.

A reducing agent (such as thioredoxin) transfers an electron to the disulfide bond, causing it to break and generating two separate cysteine residues, each with a single unpaired electron (also known as thiyl radicals).
One of the thiyl radicals is rapidly converted into a stable thiol group by reaction with a nearby protein cysteine residue, which helps to prevent unwanted reactions.

Propagation:

One of the thiyl radicals (Cys•) on the enzyme reacts with molecular oxygen to generate a peroxide intermediate (Cys-S-O-O•).
The peroxide intermediate is then rapidly converted into a tyrosyl radical (Tyr•) on a nearby tyrosine residue by an electron transfer reaction.
The tyrosyl radical is then transferred to a substrate molecule (such as a ribonucleotide diphosphate), which initiates the radical-mediated chemistry necessary for nucleotide reduction.
Overall, the generation of the free radical in RNR enzymes is a carefully orchestrated process that allows for precise control over the production of reactive species, enabling the enzyme to carry out its essential functions in DNA synthesis and repair.

Once the two cysteine amino acids in the active site of the RNR enzyme have been used in the reaction to generate the free radical, they are converted to a disulfide bond. This disulfide bond then needs to be reduced in order for the enzyme to continue functioning. In class I RNR enzymes, this reduction is accomplished by a flavoprotein known as thioredoxin reductase, which transfers electrons from NADPH to a molecule of thioredoxin. Thioredoxin then reduces the disulfide bond in the RNR enzyme's active site, regenerating the cysteine residues and allowing the enzyme to continue its catalytic cycle. In class II and III RNR enzymes, different electron transfer proteins are involved in the reduction of the disulfide bond.

This reduction leads to the formation of a thiyl radical on the cysteine residue and a transient tyrosyl radical on a nearby tyrosine residue. A thiyl radical is a highly reactive species that contains an unpaired electron on the sulfur atom of a cysteine residue. It is formed in the RNR enzyme during the process of generating the free radical required for the enzyme's catalytic activity. The thiyl radical plays a critical role in the enzyme's mechanism by abstracting a hydrogen atom from the substrate, thereby initiating the radical transfer process that leads to the generation of deoxyribonucleotides. The tyrosyl radical is then transferred to a substrate molecule, which initiates the radical-mediated chemistry necessary for nucleotide reduction. This radical transfer process is what makes RNR enzymes unique and essential for DNA synthesis and repair in all living organisms.

The active site of the enzyme contains a tyrosine residue that is used to generate a free radical on the ribonucleotide.  The process occurs in several steps:

1. A substrate (the ribonucleotide) binds to the enzyme's active site, where it is coordinated by several amino acid residues, including the tyrosine residue.
2. An adjacent cysteine residue in the active site donates an electron to the tyrosine residue, creating a tyrosyl radical.
3. The tyrosyl radical abstracts a hydrogen atom from the substrate, creating a substrate radical and regenerating the tyrosine residue.
4. The substrate radical then reacts with the thiyl radical generated from the cysteine residue in the earlier step, resulting in the formation of a new covalent bond between the ribonucleotide and the cysteine residue.
5. This reaction produces a new cysteine residue with a thiol group, and a new substrate that has been converted to its corresponding deoxyribonucleotide.
6. Overall, the tyrosine residue acts as a mediator, transferring the radical to the substrate to enable the reduction reaction to occur.

The essential players involved in the process to generate a free radical

Generating a free radical on the ribonucleotide in RNR enzymes requires: 

1. The RNR enzyme itself contains the active site responsible for the generation of the free radical.
2. A source of electrons, which reduces the disulfide bond in the enzyme's active site. In class I and II RNRs, this source is a flavoprotein that donates electrons to the enzyme. In class III RNRs, the source is a ferredoxin.
3. The substrate ribonucleotide, which is targeted by the free radical and converted into its corresponding deoxyribonucleotide form.
4. The amino acid residues in the enzyme's active site, including the cysteine and tyrosine residues, which are essential for the formation and stabilization of the free radical.

All of these components are necessary for the RNR enzyme to function properly and carry out its crucial role in DNA synthesis and repair. The RNR enzyme can be considered irreducibly complex, as it requires the coordinated and functional interaction of multiple components to generate the free radical necessary for DNA synthesis and repair. Removal or impairment of any one of these components would render the enzyme non-functional. The individual players/subunits/substrates involved in the process of generating a free radical on the ribonucleotide in RNR enzymes would have no function on their own, and they need to be integrated in the system for the enzyme to function properly. This is a key characteristic of irreducible complexity, where the individual components of a complex system are interdependent and cannot function on their own. The probability of the individual players arising through purely random, unguided processes is warranted to be considered low to the extreme, even by many scientists due to the complexity and specificity of these systems. 

Dr. Douglas Axe, a molecular biologist and director of the Biologic Institute, has written extensively on the subject of protein evolution:

"The kind of enzyme we're talking about here is mind-bogglingly complex. It's a gigantic machine. It's not just a couple of amino acids strung together. You're talking about a machine that has multiple moving parts, has different metals, it has different ligands that it has to bind to. It has to be regulated. It's an incredibly complex thing."  ( he was not referring to the ribonucleotide reductase specifically, but he was speaking more generally about the complexity of certain enzymes, including many proteins involved in cellular metabolism. ) 

Dr. Axe made this statement in a 2016 interview with The College Fix, in which he discussed his research on protein evolution and his skepticism of the idea that complex proteins like RNR enzymes could have arisen by chance through naturalistic processes.

The mechanism in Class II RNR enzymes

The Class II RNR enzyme is a homodimer, meaning it consists of two identical subunits. Each subunit contains three domains: a substrate-binding domain, a radical-generating domain, and a catalytic domain.

1. The substrate-binding domain of each subunit binds to a ribonucleotide, specifically the 2'-OH group of the ribose sugar.
2. The radical-generating domain of each subunit contains a cofactor called adenosylcobalamin (AdoCbl), which is a form of vitamin B12. The AdoCbl is converted to a highly reactive species called 5'-deoxyadenosyl radical (dAdo•) by the transfer of an electron from a nearby iron-sulfur cluster.
3. The dAdo• radical is then transferred from one subunit to the other, across the dimer interface, where it reacts with the ribonucleotide bound to the substrate-binding domain. The dAdo• radical abstracts a hydrogen atom from the 2'-OH group of the ribose sugar, generating a carbon-centered radical on the sugar ring.
4. The carbon-centered radical is then stabilized by the radical-generating domain, which donates an electron to the radical, converting it to a stable intermediate.
5. The stable intermediate is then transferred to the catalytic domain, where it undergoes a series of proton and electron transfers, leading to the reduction of the ribonucleotide to a deoxyribonucleotide.
6. Finally, the deoxyribonucleotide product is released, and the enzyme returns to its starting state, ready to bind to another ribonucleotide substrate and repeat the cycle.

The mechanism of Class II RNR enzymes is highly complex and involves multiple subunits, cofactors, and radical intermediates. 

Class II RNR enzymes differ from the other two classes of RNR enzymes (Class I and Class III) in both their structure and mechanism. The most notable structural difference is that Class II RNR enzymes are homodimers, meaning that they consist of two identical subunits, while Class I and III RNR enzymes are heterodimers, meaning they consist of two different subunits. In terms of mechanism, Class II RNR enzymes use a radical-based mechanism, while Class I and III RNR enzymes use a different mechanism that involves the formation of a free radical on a cysteine residue in the active site of the enzyme. In Class II RNR enzymes, the radical is generated on a cofactor called adenosylcobalamin (AdoCbl), while in Class I and III RNR enzymes, the radical is generated on a conserved cysteine residue. Another difference between the three classes of RNR enzymes is the way in which they are regulated. Class II RNR enzymes are typically regulated at the level of gene expression, meaning that their activity is controlled by the production or degradation of the enzyme itself. In contrast, Class I and III RNR enzymes are regulated by a variety of mechanisms, including allosteric regulation, protein-protein interactions, and post-translational modifications. While all three classes of RNR enzymes catalyze the conversion of ribonucleotides to deoxyribonucleotides, they differ in their structural features, reaction mechanisms, and modes of regulation.

The complexity of Class II RNR enzymes presents a challenge to understanding how they could have evolved from simpler precursors. One suggestion of evolutionary relatedness to the other versions is the fact that both,  Class I and Class II RNR enzymes contain an iron-sulfur cluster, and both use a radical-generating cofactor to initiate nucleotide reduction. The iron-sulfur clusters in Class I and Class II RNR enzymes are similar, but not identical. Both classes of enzymes use iron-sulfur clusters to transport electrons during the nucleotide reduction process, but the specific structures and functions of these clusters differ between the two classes.

Comparing the iron-sulfur cluster between Class I, and Class II RNR enzymes

Iron-sulfur clusters are found in a wide range of proteins in almost all forms of life, including bacteria, archaea, and eukaryotes. There are some cells or organisms that do not contain enzymes or proteins with iron-sulfur clusters, either because they do not require them for their metabolic processes or because they have evolved alternative mechanisms for performing the same functions. For example, some anaerobic bacteria can use other types of electron carriers, such as flavoproteins or quinones, instead of iron-sulfur clusters for their energy metabolism. In addition, some organisms may have evolved different mechanisms for DNA repair and other cellular processes that do not rely on iron-sulfur clusters. Iron-sulfur clusters are highly versatile and are involved in a wide range of cellular processes, and they are considered to be one of the oldest and most conserved cofactors in biology. Therefore, it is unlikely that cells or organisms could completely do without iron-sulfur clusters or an equivalent mechanism to carry out their essential metabolic processes.  The origin of iron-sulfur clusters is considered to be an origin of life problem. Iron-sulfur clusters are one of the oldest and most widespread cofactors in biology, and they are found in a wide range of proteins involved in various cellular processes, including energy metabolism, DNA replication and repair, and regulation of gene expression. 

The "iron-sulfur world" hypothesis

The "iron-sulfur world" hypothesis is a theory regarding the origin of life on Earth that suggests that life may have originated in an environment rich in iron and sulfur minerals. This hypothesis proposes that the first living organisms may have used iron-sulfur clusters as a primitive form of enzymatic activity, which could have facilitated the chemical reactions necessary for the emergence of life.

The iron-sulfur world hypothesis is based on several observations. First, iron and sulfur are abundant elements that were likely present in the early Earth's crust and oceans. Second, iron-sulfur clusters are highly versatile and can catalyze a wide range of chemical reactions, including those involved in energy metabolism, DNA replication and repair, and the synthesis of amino acids and other organic molecules. Third, iron-sulfur clusters are highly conserved in modern organisms, suggesting that they may have been present in the last universal common ancestor (LUCA) of all life forms.

According to the iron-sulfur world hypothesis, the first living organisms would have used iron-sulfur clusters to carry out primitive forms of metabolic and enzymatic activity, which could have allowed them to harness the energy and resources available in the early Earth's environment. Over time, these organisms would have generated more complex metabolic pathways and biochemical processes, leading to the emergence of the diverse forms of life that exist today. 

Iron-sulfur clusters Class I RNR enzymes

In Class I RNR enzymes, the iron-sulfur cluster is a [Fe-S] cluster that consists of two iron ions and two sulfur atoms coordinated by cysteine residues in the protein. This cluster serves as an electron carrier, transferring electrons from the radical-generating cofactor to the active site of the enzyme where nucleotide reduction occurs.

In contrast, the iron-sulfur cluster in Class II RNR enzymes is a [Fe4S4] cluster that consists of four iron ions and four sulfur atoms coordinated by cysteine residues. This cluster is also involved in electron transport during nucleotide reduction, but its structure and function differ from that of the [Fe-S] cluster in Class I RNR enzymes.

Furthermore, the biosynthesis pathways for the iron-sulfur clusters in Class I and Class II RNR enzymes are similar in some respects, but differ in others.

In Class I RNR enzymes, the [Fe-S] cluster is synthesized by a complex set of enzymes called the NifS/NifU system. This system involves the transfer of sulfur from cysteine to a scaffold protein, followed by the insertion of iron ions to form the complete cluster. The [Fe-S] cluster is then incorporated into the RNR enzyme during its maturation process.

Iron-sulfur clusters Class I RNR enzymes

In Class II RNR enzymes, the [Fe4S4] cluster is also synthesized by the NifS/NifU system, but the assembly process is more complex. In addition to the transfer of sulfur from cysteine to the scaffold protein, the assembly of the [Fe4S4] cluster requires the involvement of several accessory proteins. These proteins are thought to help with the coordination of the iron ions and the formation of the cluster structure. Once the [Fe4S4] cluster is assembled, it is incorporated into the RNR enzyme during maturation.

Overall, the biosynthesis pathways for the iron-sulfur clusters in Class I and Class II RNR enzymes are complex and involve multiple steps and protein components. While there are similarities between the two pathways, the differences in the structures of the two clusters mean that there are also significant differences in the details of their biosynthesis.

The biosynthesis of the iron-sulfur cluster in Class I RNR enzymes involves multiple enzymes and protein components. The exact number of enzymes involved can vary depending on the organism and the specific details of the biosynthetic pathway, but typically there are at least three enzymes involved in the process. The first enzyme is called NifS, which is responsible for transferring sulfur from cysteine to a specialized scaffold protein called IscU. IscU then binds iron ions and facilitates their incorporation into the growing iron-sulfur cluster. The second enzyme involved in the process is NifU, which serves as a scaffold for the assembly of the iron-sulfur cluster. NifU interacts with IscU and other proteins to coordinate the incorporation of sulfur and iron ions into the cluster. Finally, the third enzyme involved in the process is a specialized chaperone protein called HscA/HscB. This protein helps to prevent the premature aggregation of the nascent iron-sulfur cluster and ensures its proper folding and incorporation into the RNR enzyme. Other proteins may also be involved in the biosynthesis of the Class I RNR iron-sulfur cluster, and the specific details of the process can vary depending on the organism and environmental conditions. The simplest biosynthesis pathway for the Class I RNR iron-sulfur cluster involves two enzymes: NifS and IscA. NifS transfers sulfur to IscA, which then binds iron ions to form the [Fe-S] cluster. The [Fe-S] cluster is then incorporated into the RNR enzyme during its maturation process.

The biosynthesis of the iron-sulfur cluster in Class II RNR enzymes involves more players than in Class I RNR enzymes. The exact number of players involved can vary depending on the organism and the specific details of the biosynthetic pathway, but typically there are at least six proteins involved in the process.

The first enzyme involved in the process is NifS, which transfers sulfur to a protein called IscA.
The second enzyme is called SufB, which interacts with IscA to assemble a [2Fe-2S] cluster.
The third enzyme is called SufC, which binds the [2Fe-2S] cluster and then interacts with SufB to assemble a [4Fe-4S] cluster.
The fourth enzyme is called SufD, which binds the [4Fe-4S] cluster and then interacts with SufC to facilitate its transfer to the RNR enzyme.
The fifth protein involved in the process is called SufA, which helps to transfer the [4Fe-4S] cluster from SufD to the RNR enzyme.
Finally, a sixth protein called SufE has also been implicated in the process, although its exact role is not yet fully understood.

The biosynthesis pathway for the iron-sulfur cluster in Class II RNR enzymes is complex and involves multiple steps and protein components. The involvement of multiple proteins in the process likely reflects the greater complexity of the [4Fe-4S] cluster itself.

Quality control in producing the iron-sulfur clusters

RNR enzymes contain Fe-S clusters, and errors in Fe-S cluster synthesis or assembly can lead to enzyme dysfunction and impaired DNA synthesis. Therefore, the error check process for Fe-S cluster synthesis in RNR enzymes is critical for maintaining proper enzyme function.

One of the ways that cells prevent errors in Fe-S cluster synthesis in RNR enzymes is through a protein called NrdH-redoxin, which serves as a chaperone for the Fe-S clusters during their assembly. NrdH-redoxin helps to prevent misincorporation of iron or sulfur atoms into the Fe-S cluster, which could result in impaired enzyme function.

Another mechanism for error-checking Fe-S cluster synthesis in RNR enzymes is through the use of iron-responsive element (IRE) sequences in the messenger RNA (mRNA) that encodes the RNR enzyme. IRE sequences are recognized by iron regulatory proteins (IRPs), which can bind to the mRNA and regulate its translation into protein. If the cell detects an error in Fe-S cluster synthesis, IRPs can block translation of the mRNA, preventing the synthesis of defective RNR enzymes.

Finally, cells can also use quality control mechanisms to monitor the activity of RNR enzymes with Fe-S clusters. For example, cells may increase the expression of other Fe-S cluster-containing proteins to compensate for impaired RNR enzyme function, or they may activate stress response pathways to help the cell cope with the effects of defective Fe-S clusters.

Another mechanism for error-checking Fe-S cluster synthesis in RNR enzymes is through the use of iron-responsive element (IRE) sequences in the messenger RNA (mRNA) that encodes the RNR enzyme. IRE sequences are recognized by iron regulatory proteins (IRPs), which can bind to the mRNA and regulate its translation into protein. If the cell detects an error in Fe-S cluster synthesis, IRPs can block translation of the mRNA, preventing the synthesis of defective RNR enzymes.

Cells can also use quality control mechanisms to monitor the activity of RNR enzymes with Fe-S clusters. For example, cells may increase the expression of other Fe-S cluster-containing proteins to compensate for impaired RNR enzyme function, or they may activate stress response pathways to help the cell cope with the effects of defective Fe-S clusters.

In both Class I and Class II RNR enzymes, the quality control mechanism involves as well a protein called SufBCD, which recognizes and degrades iron-sulfur clusters that are improperly assembled or damaged. SufBCD acts as a "proofreading" mechanism, checking the quality of the iron-sulfur cluster before it is incorporated into the RNR enzyme. If the cluster fails the quality control check, it is disassembled and its components are recycled to prevent their incorporation into the RNR enzyme.

Overall, the quality control mechanism in the biosynthesis of iron-sulfur clusters in RNR enzymes is an important safeguard that helps to ensure the proper function of these enzymes in maintaining genome integrity and preventing DNA damage.

The proper synthesis and function of RNR enzymes, as well as many other biological molecules and systems, requires in most, if not in all cases, the implementation of error check and repair systems from the start. Without these systems, the error rate would likely be too high for the enzyme or system to function properly, potentially leading to cellular damage, disease, or even death. The existence of error check and repair systems are often interdependent with other biological processes, such as the synthesis and function of RNR enzymes, DNA replication and repair, and many other processes. In many cases, the proper functioning of these biological processes relies on the presence of error check and repair systems to maintain their integrity and prevent damage or errors from occurring. This interdependence between different biological processes is a common feature of living organisms and is often powerful evidence for the necessity of a mind with foreknowledge and foresight to instantiate such complexity and sophistication in biological systems.

SufBCD, the error check and repair machine in the cell

SufBCD is a protein complex involved in the biogenesis of iron-sulfur clusters, which are important cofactors found in a wide range of proteins involved in various cellular processes, including electron transport, DNA replication and repair, and regulation of gene expression. The SufBCD complex is composed of three subunits: SufB, SufC, and SufD. SufB is a peripheral membrane protein that interacts with the inner membrane of bacteria, while SufC and SufD are cytoplasmic proteins. SufB contains a conserved domain that is involved in binding iron-sulfur clusters, while SufC and SufD interact with each other to form a nucleotide-binding domain that binds ATP and helps to regulate the activity of the complex. The complex also interacts with other proteins involved in iron-sulfur cluster biogenesis, such as SufA and SufE, to facilitate the transfer and incorporation of the iron-sulfur clusters into target proteins.

The SufBCD complex is important for the survival of bacteria in environments with limited iron availability, as iron-sulfur clusters are necessary for the activity of many essential enzymes involved in metabolism and other cellular processes. Dysfunction or deficiency of the SufBCD complex can lead to impaired iron-sulfur cluster biogenesis and various cellular defects, including sensitivity to oxidative stress, DNA damage, and antibiotic treatments. The SufBCD complex is composed of three subunits: SufB, SufC, and SufD. SufB is a peripheral membrane protein that interacts with the inner membrane of bacteria, while SufC and SufD are cytoplasmic proteins.

SufB contains a conserved domain that is involved in binding iron-sulfur clusters, while SufC and SufD interact with each other to form a nucleotide-binding domain that binds ATP and helps to regulate the activity of the complex.
The action core of the SufBCD complex is the SufB subunit, which contains a conserved domain that is involved in the binding of iron-sulfur clusters. This domain is known as the Fe-S cluster-binding domain, and it contains three cysteine residues that are involved in coordinating the binding of the iron and sulfur ions that make up the cluster. The Fe-S cluster-binding domain is located near the N-terminus of the SufB protein and is essential for the function of the complex. The co-factor of the SufBCD complex is the iron-sulfur cluster, which is a small, inorganic molecule composed of iron and sulfur ions. Iron-sulfur clusters are essential cofactors found in many proteins involved in cellular processes, including energy metabolism, DNA replication and repair, and regulation of gene expression. They are known for their ability to transfer electrons and to act as redox centers in many enzymatic reactions.
The biosynthesis of iron-sulfur clusters occurs through a complex pathway that involves several proteins, including the SufBCD complex. In this pathway, sulfur and iron ions are imported into the cell through various transport systems and are assembled into iron-sulfur clusters by the action of specific proteins. The SufBCD complex is involved in the later stages of this pathway, where it helps to transfer the iron-sulfur clusters to target proteins, where they can be incorporated into the active sites of enzymes. The SufBCD complex is an essential component of the iron-sulfur cluster biosynthesis pathway, and plays a critical role in the assembly and transfer of iron-sulfur clusters to target proteins.

The synthesis pathway to make SufBCD

The SufBCD enzyme is composed of three different subunits: SufB, SufC, and SufD. The biosynthesis pathway of the SufBCD enzyme involves the coordinated expression and assembly of these subunits, as well as the synthesis and incorporation of the iron-sulfur clusters that are required for its function. Here is an overview of the synthesis pathway of the SufBCD enzyme:

1. Transcription: The genes encoding the SufBCD subunits are transcribed from the DNA into messenger RNA (mRNA) by the RNA polymerase enzyme.
2. Translation: The mRNA is then translated into protein by ribosomes, with each subunit being synthesized separately.
3. Chaperone proteins: As the individual subunits are synthesized, they are bound and stabilized by chaperone proteins, which prevent them from aggregating and ensure proper folding.
4. Assembly: Once all three subunits have been synthesized and properly folded, they are assembled into the SufBCD enzyme complex. This assembly process is coordinated by a series of accessory proteins, which help to ensure that the subunits are correctly positioned and oriented relative to each other.

Iron-sulfur cluster synthesis: The final step in the biosynthesis of the SufBCD enzyme involves the synthesis and incorporation of the iron-sulfur clusters that are required for its function. This process is mediated by a separate set of accessory proteins, which help to guide the assembly of the clusters and ensure that they are properly incorporated into the subunits of the enzyme. Overall, the biosynthesis pathway of the SufBCD enzyme is a complex and tightly regulated process that involves the coordinated expression, folding, and assembly of multiple protein subunits, as well as the synthesis and incorporation of the iron-sulfur clusters that are required for its function.

The accessory proteins involved in the synthesis of the Iron-sulfur cluster used in SufBCD

The biosynthesis of iron-sulfur clusters and their incorporation into proteins like SufBCD is a complex process that involves the coordination of multiple accessory proteins. Here is an overview of some of the accessory proteins involved in this process:

1. SufA: This protein is an iron-sulfur cluster scaffold protein that helps to assemble the iron-sulfur clusters that are required for the function of the SufBCD enzyme. SufA binds to the iron and sulfur atoms and helps to coordinate their assembly into a stable cluster.
2. SufE: This protein is an iron-sulfur cluster carrier protein that helps to deliver the clusters from the SufA scaffold protein to the target proteins like SufBCD. SufE binds to the clusters and then interacts with other proteins to facilitate their transfer.
3. SufB: This is one of the subunits of the SufBCD enzyme itself, but it also functions as an accessory protein during the biosynthesis of iron-sulfur clusters. SufB helps to deliver iron and sulfur to the SufA scaffold protein, and also interacts with SufC and SufD to coordinate the assembly of the iron-sulfur clusters within the SufBCD complex.
4. SufC: This is another subunit of the SufBCD enzyme, and it plays a critical role in the biosynthesis of iron-sulfur clusters by interacting with both SufB and SufD to coordinate the assembly of the clusters within the enzyme complex.

Overall, the biosynthesis of iron-sulfur clusters and their incorporation into proteins like SufBCD is a complex process that requires the coordination of multiple accessory proteins. These accessory proteins help to ensure that the clusters are assembled and delivered to their target proteins in a precise and controlled manner, which is essential for the proper function of the enzyme.

Another chicken & egg problem

There is another chicken-and-egg problem when it comes to the biosynthesis of iron-sulfur clusters since many enzymes that are involved in the synthesis and incorporation of these clusters themselves require iron-sulfur clusters for their function. Several possible solutions to this problem have been proposed. One is that the first iron-sulfur clusters were synthesized by non-enzymatic processes, such as the reaction of iron and sulfur in the presence of a reducing agent. The non-enzymatic synthesis of iron-sulfur clusters by simple chemical reactions would likely be a very unspecific and inefficient process. Some researchers have proposed that early earth environments, such as hydrothermal vents, would have provided conditions that were conducive to the formation of iron-sulfur clusters through specific mineral catalysis or other mechanisms. These environments would have also provided a source of reducing agents or other reactants that could have facilitated the formation of these clusters in a more controlled manner. While hydrothermal vents and other early earth environments could have provided specific conditions that favored the formation of iron-sulfur clusters, the overall non-specificity of the process would however still be a challenge that cannot be overstated enough. Another claim is that the earliest enzymes that required iron-sulfur clusters for their function were simpler and more primitive than modern enzymes, and were able to operate with simpler or more easily assembled clusters. Over time, these enzymes could have progressed to become more complex and sophisticated and could have developed the ability to synthesize and incorporate more complex iron-sulfur clusters. There is however a large gap between the simpler, more primitive enzymes that would have operated with simpler iron-sulfur clusters, and the highly regulated, multi-step processes involving multiple enzymes that are required for the biosynthesis of iron-sulfur clusters in modern cells. Another hypothesis states that many of the enzymes involved in modern iron-sulfur cluster biosynthesis would have evolved from more ancient enzymes that performed related functions in early life forms. But life was not even extant at such a stage. This is just one tiny problem, among many other factors and processes involved in the emergence of life on Earth, which is unsolved, equally to this problem,  and the formation of iron-sulfur clusters is just one piece of a larger puzzle.


The mechanism in Class III RNR enzymes

The reaction mechanism of Class III RNR enzymes can be divided into two stages: initiation and propagation.

Initiation:

The reaction starts with the binding of a substrate, which is a ribonucleotide, to the enzyme's active site.
The substrate is then converted to a radical species by a radical-generating cofactor, such as adenosylcobalamin or glycyl radical, that is associated with the enzyme.
The radical generated on the substrate abstracts a hydrogen atom from a nearby cysteine residue on the enzyme, forming a cysteine radical and a substrate radical.

Propagation:

The substrate radical then undergoes a series of electron and proton transfers within the active site, which leads to the reduction of the substrate to its corresponding deoxyribonucleotide.

The electron and proton transfers involve the participation of several amino acid residues and cofactors that are located within the enzyme's active site.
The cysteine radical generated in the initiation stage is then regenerated back to its original form by a reducing agent, such as thioredoxin or glutaredoxin.

Class III ribonucleotide reductase (RNR) enzymes do not contain iron-sulfur clusters.

Iron-sulfur clusters are cofactors found in Class I and Class II RNR enzymes, which use a different mechanism to catalyze the reduction of ribonucleotides to deoxyribonucleotides. Class III RNR enzymes, on the other hand, use a radical mechanism that involves the participation of radical-generating cofactors, such as adenosylcobalamin or glycyl radical.

It should be noted that not all RNR enzymes contain iron-sulfur clusters. In addition to Class I RNR enzymes, some Class II RNR enzymes also contain iron-sulfur clusters, while others use different cofactors, such as flavodoxin or heme, to generate radicals.

The active site of Class III ribonucleotide reductase (RNR) enzymes contains several key components that are involved in the reduction of ribonucleotides to deoxyribonucleotides. The following steps outline the general process that occurs in the active site of Class III RNR enzymes:

1. Substrate binding: The ribonucleotide substrate binds to the active site of the enzyme, where it interacts with a radical-generating cofactor, such as adenosylcobalamin or glycyl radical. This interaction results in the generation of a radical on the substrate.
2. Radical transfer: The radical on the substrate undergoes a radical transfer reaction, where it is transferred to a nearby cysteine residue on the enzyme. This forms a cysteine radical and a substrate radical.
3. Propagation: The substrate radical undergoes a series of electron and proton transfers within the active site, which leads to the reduction of the substrate to its corresponding deoxyribonucleotide. This process involves the participation of several amino acid residues and cofactors within the active site, which help to facilitate the transfer of electrons and protons.
4. Cysteine regeneration: The cysteine radical generated in step 2 is then regenerated back to its original form by a reducing agent, such as thioredoxin or glutaredoxin. This allows the enzyme to continue to catalyze the reduction of additional ribonucleotides.

Overall, the active site of Class III RNR enzymes plays a critical role in facilitating the radical mechanism used to reduce ribonucleotides to deoxyribonucleotides. The active site contains specific amino acid residues and cofactors that help to generate and transfer radicals, as well as facilitate the electron and proton transfers necessary for catalysis

Adenosylcobalamin

Adenosylcobalamin has one of the most complex structures among all vitamins. It consists of a large, complex molecule known as a corrin ring, which is bound to a central cobalt ion. Cobalt ion is a positively charged ion of the element cobalt. Cobalt is a transition metal that can form different ions depending on its oxidation state. In its ionic form, cobalt can have a +2 or +3 charge. In adenosylcobalamin, the cobalt ion is in the +3 oxidation state and is bound to the corrin ring, forming the core of the coenzyme. The corrin ring is a large, complex organic molecule that is the central part of the adenosylcobalamin coenzyme. It consists of a planar tetrapyrrole ring that contains four nitrogen atoms and is similar in structure to the heme group found in hemoglobin. However, the corrin ring is larger and more complex than the heme group.

The corrin ring has a unique three-dimensional structure that allows it to bind to the cobalt ion at its center and to interact with other molecules during biochemical reactions. It also has several functional groups, including carboxyl and methyl groups, that play important roles in the chemistry of the coenzyme. The corrin ring is synthesized by certain bacteria and archaea, and it is not produced by humans or other animals. It is an essential component of the adenosylcobalamin coenzyme, which is required for several important metabolic processes in the body, including the breakdown of fatty acids and amino acids. Attached to one end of the corrin ring is a nucleotide called adenosine, which serves as a binding site for the enzyme that uses the coenzyme. The other end of the corrin ring is the site where the reaction takes place. The complex structure of adenosylcobalamin is necessary for its ability to serve as a cofactor in a wide range of enzymatic reactions in the body. Its unique structure allows it to act as a "molecular carrier" that can shuttle groups of atoms between different molecules during metabolic processes.

Adenosylcobalamin, also known as coenzyme B12, is a coenzyme that is involved in several important enzymatic reactions in the body. It is a type of cobalamin, which is a group of compounds that contain the metal ion cobalt. Adenosylcobalamin is important for the metabolism of certain amino acids, as well as the breakdown of fatty acids and the synthesis of certain neurotransmitters. It is also involved in the production of energy from glucose through a process called the Krebs cycle. The molecule itself consists of a corrin ring that is coordinated to the cobalt ion. Attached to one end of the corrin ring is a nucleotide called adenosine, which serves as a binding site for the enzyme that uses the coenzyme. The other end of the corrin ring is the site where the reaction takes place. Adenosylcobalamin is produced in the body through a complex biosynthesis pathway that involves several enzymes and cofactors. In this process, cobalt is incorporated into a precursor molecule, which is then modified and processed into the final adenosylcobalamin molecule. Deficiencies in the enzymes involved in this pathway can lead to a range of health problems, including anemia and neurological disorders.

The biosynthesis steps of Adenosylcobalamin

The biosynthesis of Adenosylcobalamin (AdoCbl) involves more than 30 enzymatic steps, and it occurs mainly in certain bacteria and archaea. Here is a brief overview of the biosynthesis steps:

1. Synthesis of uroporphyrinogen III: This step involves the conversion of glutamate to uroporphyrinogen III, which is a precursor of the corrin ring.
2. Formation of the corrin ring: The corrin ring is formed from the condensation of four molecules of uroporphyrinogen III, followed by the insertion of a cobalt ion into the center of the ring.
3. Reduction of the cobalt ion: The cobalt ion in the corrin ring is reduced to cobalt(I) by a series of enzymes.
4. Attachment of aminopropanol: The cobalt(I) ion is then ligated by an aminopropanol molecule, forming cobalt-precorrin-4.
5. Methylation: A series of enzymes catalyze the addition of several methyl groups to cobalt-precorrin-4, resulting in the formation of cobalt-precorrin-5.
6. Formation of the nucleotide loop: A nucleotide loop is added to cobalt-precorrin-5 to form cobalt-precorrin-6A.
7. Attachment of the nucleotide loop: The nucleotide loop is transferred from cobalt-precorrin-6A to cob(I)alamin, forming Adenosylcobalamin (AdoCbl).

The biosynthesis of AdoCbl is a complex process that involves the coordination of multiple enzymes and cofactors. It requires the input of several different metabolic pathways, including those involved in the synthesis of amino acids, nucleotides, and porphyrins.

The biosynthesis of iron-sulfur clusters and the assembly of the RNR Class III enzyme are both complex and highly regulated processes that require multiple enzymes and cofactors. It is unlikely that these processes could have arisen spontaneously in a prebiotic environment.

What-when-how: All ribonucleotide reduction proceeds via controlled free-radical-based chemistry, in which a free radical amino acid residue of an RNR generates a substrate radical by abstracting a hydrogen atom from C3′ of the substrate, to facilitate the leaving of the OH group on the vicinal C2′. A thiyl group of a cysteine residue performs this function. Two additional redox-active cysteine residues then provide the reducing equivalents for the subsequent reduction at C2′. This general mechanism has strong experimental support for class I and II enzymes. The three-dimensional structure of the catalytic site of the E. coli class I enzyme beautifully fits this mechanism. For class III RNR, the evidence for a similar mechanism is indirect.7


Getting the right balance of nucleotides

Anne Trafton, MIT News (2016): Cell survival depends on having a plentiful and balanced pool of the four chemical building blocks that make up DNA,  A, G, C, and T. However, if too many of these components pile up, or if their usual ratio is disrupted, that can be deadly for the cell. Ribonucleotide reductase (RNR) generates all four of these building blocks and maintains the correct balance among them. Unlike RNR, most enzymes specialize in converting just one type of molecule to another. “Ribonucleotide reductase is very unusual. Its fascinating that this enzyme’s active site can be molded into four different shapes.”  RNR’s interactions with its downstream products via a special effector site causes the enzyme to change its shape, determining which of the four DNA building blocks it will generate. While many other enzymes are controlled by effectors, this type of regulation usually turns enzyme activity up or down. “I can’t think of any other examples of effector binding changing what the substrate is. This is just very unusual,” Drennan says. Deoxyribonucleotides are generated from ribonucleotides, which are the building blocks for RNAs — molecules that perform many important roles in gene expression.  “There’s no other enzyme that really can do that chemistry,” she says. “It’s the only one, and it’s very different than most enzymes and has a lot of really unusual features.” RNR can take on different shapes. The enzyme’s active site — the region that binds the substrate — changes shape depending on which effector molecule is bound to a distant site on the enzyme. For this enzyme, the effector molecules are deoxynucleoside trisphosphates such as deoxyadenosine triphosphate (dATP) or thymidine triphosphate (TTP). Depending on which of these effectors is bound to the distant regulatory site, the active site can accommodate one of the four ribounucleotide substrates. Effector binding promotes the closing of part of the protein over the active site like a latch to lock in the substrate. If the wrong base is in the active site, the latch can’t close and the substrate will diffuse out. “It’s exquisitely designed so that if you have the wrong substrate in there, you can’t close up the active site,” Drennan says. “It’s a really elegant set of movements that allows for this kind of molecular screening process.” The effectors can also shut off production completely, by binding to a completely different site on the enzyme, if the pool of building blocks is getting too big. 16

Pär Nordlund (2006): An intricate interplay between gene activation, enzyme inhibition, and protein degradation regulates, together with the allosteric effects, enzyme activity and provides the appropriate amount of deoxynucleotides for DNA replication and repair. 12

Evolutionnews (2016):  It’s like a surgical robot that has a clamp with an on-off switch. The switch (the effector) turns the machine on, opening up the distant active site and letting the appropriate substrate in. The enzyme then clamps down on the substrate and “reduces” it by replacing the oxygen radical with a hydrogen. When released, the DNA building block is ready for use, the effector switches the machine off, and the enzyme is ready for the next operation. Somehow, when there are too many building blocks floating around in the cell, an effector binds to a different active site, disabling the machine. It’s uncanny how each part seems to know what’s needed and how to provide it. This involves feedback from the nucleus, where genes respond to the supply by either locking the RNR enzymes or making more of them.16

Raleigh McElvery, MIT News (2020): Many believed the enzyme, ribonucleotide reductase (RNR’s) two subunits came together and fit with perfect symmetry, like a key into a lock. “For 30 years, that’s what we thought,” says Catherine Drennan, an MIT professor of chemistry and biology and a Howard Hughes Medical Institute investigator. “But now, we can see the movement is much more elegant. The enzyme is actually performing a ‘molecular square dance,’ where different parts of the protein hook onto and swing around other parts. It’s really quite beautiful.” The combination of new techniques allowed to visualize the complex molecular dance that allows the enzyme to transport the catalytic “firepower” from one subunit to the next, in order to generate DNA building blocks. This firepower is derived from a highly reactive unpaired electron (a radical), which must be carefully controlled to prevent damage to the enzyme. According to Drennan, the team “wanted to see how RNR does the equivalent of playing with fire without getting burned.” Although this molecular dance brings the subunits together, there is still considerable distance between them: The radical must travel 35-40 angstroms from the first subunit to the second. This journey is roughly 10 times farther than the average radical transfer. The radical must then travel back to its starting place and be stored safely, all within a fraction of a second before the enzyme returns to its normal conformation.14

This is the paper that McElvery is referencing. Here are the note-worthy concluding remarks of the paper: G. Kang (2020): The elegance of this molecular design has not escaped our attention, with the one binding/catalytic step triggering the next, guiding this enzyme through a series of elaborate conformational rearrangements that produce the requisite deoxynucleotide levels for DNA biosynthesis and repair.15

E. coli Ribonucleotide Reductase Has Three Different Nucleotide-Binding Sites

R. H. Garrett (2008): The enzyme system for deoxynucleotide diphosphate (dNDP) formation consists of four proteins, two of which constitute the ribonucleotide reductase proper. The other two proteins, thioredoxin and thioredoxin reductase, function in the delivery of reducing equivalents (energy). The two proteins of ribonucleotide reductase are designated R1 and R2, and each is a homodimer in the holoenzyme (Figure below). 



The R1 homodimer carries two types of regulatory sites in addition to the catalytic site (the active site). Substrates bind at the catalytic site. One regulatory site—the substrate specificity site—binds various nucleotides, like ATP, dATP, dGTP, or dTTP, and which of these nucleotides is bound there determines which nucleoside diphosphate is bound at the catalytic site. The other regulatory site, the overall activity site, binds either the activator ATP or the negative effector dATP; the nucleotide bound here determines whether the enzyme is active or inactive. The 2 Fe atoms within the single active site formed by the R2 homodimer generate the free radical required for ribonucleotide reduction on a specific R2 residue, Tyr122, which in turn generates a thiyl free radical. Cys439-S initiates ribonucleotide reduction by abstracting the 3-H from the ribose ring of the nucleoside diphosphate substrate and forming a free radical on C-3. Subsequent dehydration forms the deoxyribonucleotide product.23


Catherine L Drennan (2016): RNR uses multiple allosteric mechanisms to maintain the balanced deoxyribonucleoside triphosphate (dNTP) pools that are required for accurate DNA replication. First, allosteric activity regulation modulates the overall size of deoxyribonucleotide triphosphate (dNTP) pools. ATP or dATP binding at an allosteric activity site leads to up-regulation or down-regulation of enzyme activity, respectively. In E. coli class Ia RNR, this regulation is achieved by changes in the oligomeric arrangement of the α2 and β2 subunits. When ATP is bound at the activity site, an α2β2 complex is favored. This active α2β2 complex is capable of a long-range proton-coupled electron transfer from β2 to α2, forming a transient thiyl radical on Cys439 to initiate catalysis. Alternatively, when concentrations of dATP become too high in the cell, dATP binds at the allosteric activity site and formation of an α4β4 complex is promoted. The structure of this complex was recently solved, revealing a ring of alternating α2 and β2 units that cannot form a productive electron transfer path, thus inhibiting the enzyme.

RNR communication between the allosteric site, and the active site

Ribonucleotide reductase (RNR) is an essential enzyme that plays a critical role in the synthesis of deoxyribonucleotides, the building blocks of DNA. RNRs have an allosteric regulation mechanism that allows them to regulate their activity based on the intracellular concentration of deoxyribonucleotides. The allosteric site of RNR contains a binding site for ATP or dATP, which acts as a feedback inhibitor of RNR activity. When the intracellular concentration of dATP is high, it binds to the allosteric site and inhibits RNR activity to prevent the overproduction of deoxyribonucleotides. The binding of ATP or dATP to the allosteric site induces conformational changes in the RNR enzyme, which are transmitted to the active site. This leads to the formation of an inhibitory complex at the active site that blocks the activity of RNR.

Conversely, when the intracellular concentration of deoxyribonucleotides is low, the allosteric site of RNR binds to ATP instead of dATP, which activates the enzyme. This activation occurs through a mechanism called substrate specificity modulation, in which ATP binding induces conformational changes that favor the binding of substrates to the active site. Thus, the communication between the allosteric and active sites of RNR is crucial for regulating the synthesis of deoxyribonucleotides and maintaining the appropriate balance of nucleotides in the cell.

The regulation of RNR activity through communication between the allosteric and active sites is essential for the survival of the cell, as it allows the cell to balance its production of deoxyribonucleotides with its demand for DNA synthesis. If this regulation were to be interrupted, it could lead to uncontrolled or insufficient production of deoxyribonucleotides, which could have negative consequences for DNA synthesis and ultimately for cell survival. If the regulation of RNR communication between the allosteric site and the active site is interrupted, it can lead to imbalanced dNTP pools and cellular damage. For example, in some cancer cells, mutations in the genes that encode RNR regulatory proteins have been observed, resulting in uncontrolled DNA replication and increased genomic instability. Additionally, drugs that target RNR have been developed as cancer therapeutics because they can disrupt this regulation and selectively kill rapidly dividing cancer cells by causing imbalanced dNTP pools and DNA damage.

There is a unifying mechanism for substrate specificity regulation in the most studied RNR, the E. coli class Ia enzyme. Our structures show how each specificity effector is read out at a distal allosteric site and how that information is communicated to the active site where residues rearrange such that specific hydrogen bonds can be formed with the cognate substrate base. When an effector/substrate match is discovered, the barrel is clamped and latched in preparation for catalysis. Just as DNA replication and transcription take advantage of the unique hydrogen-bonding properties of each nucleotide base, enzymatic ribonucleotide reduction also employs these unique hydrogen-bonding properties for specificity regulation. Through an elegant set of protein rearrangements, E. coli RNR screens and selects its substrate from the four potential NDPs, ensuring appropriate pools of deoxynucleotides are available for DNA biosynthesis and repair.18

The enzyme consists of four proteins, two of which constitute the ribonucleotide reductase proper. The other two proteins, thioredoxin and thioredoxin reductase, function in the delivery of reducing equivalents. ( basically the energy)
The R1 homodimer carries two types of regulatory sites in addition to the catalytic site (the active site). Substrates (ADP, CDP, GDP, and UDP) bind at the catalytic site. One regulatory site—the substrate specificity site—binds ATP, dATP, dGTP, or dTTP, and which of these nucleotides is bound there determines which nucleoside diphosphate is bound at the catalytic site. The other regulatory site, the overall activity site, binds either the activator ATP or the negative effector dATP; the nucleotide bound here determines whether the enzyme is active or inactive.

Overall assembly of RNR enzymes

The overall assembly process of RNR enzymes can be divided into several steps:

1. Synthesis of subunits: The subunits of RNR enzymes are synthesized by the ribosomes and are either encoded by the same gene (class III RNR) or by multiple genes (class I and II RNR).
2. Formation of dimeric or multimeric subunit complexes: The subunits of RNR enzymes associate to form dimeric or multimeric subunit complexes. In class I RNR enzymes, the large α subunit forms a dimer with the small β subunit. In class II RNR enzymes, the α and β subunits form a heterodimeric complex. In class III RNR enzymes, a single polypeptide chain forms the active enzyme.
3. Binding of cofactors: RNR enzymes require cofactors, such as iron-sulfur clusters, radicals, and ATP, for their activity. These cofactors bind to specific domains or motifs on the subunits and are critical for the proper function of the enzyme.
4. Allosteric regulation: RNR enzymes are allosterically regulated by various effectors that can activate or inhibit enzyme activity. These effectors bind to specific sites on the enzyme and cause conformational changes that affect enzyme activity.
5. Assembly of the holoenzyme: The dimeric or multimeric subunit complexes with bound cofactors and allosteric effectors associate to form the active holoenzyme complex.

The assembly process of RNR enzymes is a complex and coordinated process that involves the synthesis of subunits, the formation of subunit complexes, the binding of cofactors, and the allosteric regulation of enzyme activity. The proper assembly of RNR enzymes is essential for the de novo synthesis of deoxyribonucleotides and DNA replication.

The binding of cofactors to specific domains or motifs on RNR enzymes is determined by the amino acid sequence and structural features of these domains or motifs. For example, iron-sulfur clusters typically bind to specific cysteine residues that are coordinated by the iron atoms of the cluster. The amino acid sequence and structural context of these cysteine residues help to determine the specificity of iron-sulfur cluster binding. The amino acid sequence and structural context of the cysteine residues in the RNR enzyme determine the specificity of iron-sulfur cluster binding. If the cysteine residues are not present or are not in the proper sequence or structural context, the iron-sulfur cluster would not be able to bind to the enzyme. Therefore, the amino acid sequence and structure of the RNR enzyme must be precisely maintained to ensure the proper binding of iron-sulfur clusters and the formation of an active holoenzyme complex. The precise amino acid sequence and structural context required for cofactor binding and enzyme function present significant challenges to the step-wise origin and evolutionary development of enzymes like RNR. It is challenging to imagine how a protein with no cofactor binding site could evolve to bind a cofactor in a specific location with high specificity. Evolutionary intermediates would likely have reduced enzymatic activity or could be detrimental to the organism's survival if they bind to the wrong cofactor or bind the cofactor in the wrong location.

One claimed possibility is that the ancestral protein would have had weak, promiscuous binding to a range of cofactors, with further evolution leading to the specificity observed in modern enzymes. Another possibility is that the binding sites and cofactor specificity could have evolved in parallel with the evolution of the protein's catalytic activity. Similarly, radicals, such as the glycyl radical in class III RNR enzymes, are generated by radical SAM enzymes that recognize specific sequence motifs and catalyze radical formation through a conserved mechanism. A promiscuous binding site would however not be sufficient for proper enzymatic function. Enzymes often require precise binding of specific cofactors to perform their biological functions effectively. Weak or non-specific binding may lead to low enzyme activity or binding to non-native cofactors that can interfere with proper enzyme function. RNR enzymes require precise binding of multiple cofactors, and the step-wise evolution of the required cofactor binding sites and specificity presents significant challenges.

ATP and other nucleotides bind to specific binding pockets on RNR enzymes that have complementary amino acid sequences and structural features. These binding pockets are typically located in specific domains of the enzyme that interact with the nucleotide cofactor. Allosteric effectors, such as nucleotide triphosphates or dATP, also bind to specific sites on RNR enzymes, causing conformational changes that affect enzyme activity. These sites are typically located in specific domains or motifs that have evolved to interact with the allosteric effector. A stepwise evolution of the nucleotide-binding pockets and allosteric effector sites in RNR enzymes would also face significant challenges, similar to those faced by the evolution of cofactor binding sites. The precise amino acid sequence and structural context required for specific nucleotide binding and allosteric regulation also present significant challenges for step-wise evolution. In the absence of these features, the binding of nucleotides or allosteric effectors may be non-specific or non-functional, leading to low enzymatic activity or even detrimental effects on the organism's survival.

Therefore, the step-wise emergence of RNR enzymes by multiple mutations coordinating the evolution of multiple domains or motifs to create the specific binding sites required for nucleotide binding and allosteric regulation would have been in the realm of the impossible. In special in face of the fact that intermediate stages would have resulted in non-functional enzymes.

The assembly of the holoenzyme

The assembly of the holoenzyme involves multiple steps that occur sequentially:

1. Synthesis of the individual subunits: Each subunit of the holoenzyme is synthesized separately.
2. Folding of the individual subunits: Each subunit must fold into its native conformation.
3. Binding of cofactors: The subunits bind cofactors required for the enzyme activity.
4. Formation of subunit complexes: Dimeric or multimeric subunit complexes are formed, which may involve interactions between the subunits.
5. Binding of allosteric effectors: Allosteric effectors bind to specific sites on the enzyme, causing conformational changes that affect enzyme activity.
6. Association of subunit complexes: The subunit complexes associate to form the active holoenzyme complex.

These steps occur in a specific order, with each step building upon the previous one. For example, the folding of the individual subunits needs to be completed before the subunit complexes can be formed. Similarly, the binding of cofactors may need to occur before allosteric effectors can bind to the enzyme. The final step of association of the subunit complexes to form the holoenzyme complex requires multiple subunits to come together in a specific orientation, which is essential for enzyme activity.

The process of assembly of the holoenzyme is coordinated through various mechanisms that ensure the correct order of steps and proper folding and assembly of each subunit. These mechanisms include chaperones, protein-protein interactions, and the binding of cofactors and allosteric effectors. Chaperones are specialized proteins that facilitate the proper folding of newly synthesized proteins by preventing them from forming incorrect conformations. In the case of the holoenzyme, chaperones may help the individual subunits fold correctly before they are assembled into complexes. Protein-protein interactions between the subunits also play a critical role in ensuring that the correct subunits come together to form the complex. The binding of cofactors and allosteric effectors also helps to coordinate the process. Cofactors are required for the enzyme activity and must be bound to their respective subunits before the subunits can be assembled into complexes. Similarly, the binding of allosteric effectors to the enzyme triggers conformational changes that may be necessary for the subunits to properly associate and form the holoenzyme complex. The process of assembly of the holoenzyme is a highly coordinated and regulated process that relies on multiple mechanisms to ensure the correct order of steps and proper folding and assembly of each subunit.

Monitoring of the assembly process, and repair mechanisms

There is evidence that various chaperones and co-chaperones, which are proteins that assist in the folding and assembly of other proteins, are involved in the process. For example, the heat shock protein 90 (Hsp90) has been shown to interact with the RNR subunit RRM2 and assist in its folding and assembly into the active holoenzyme complex. Other proteins, such as the cochaperone p23 and the Hsp70 family member Hsc70, have also been implicated in the assembly of RNR enzymes. Additionally, there is evidence that post-translational modifications, such as phosphorylation and acetylation, may also play a role in regulating the assembly of RNR enzymes. When errors in the assembly of RNR enzymes are detected, quality control mechanisms are activated to prevent the release of improperly folded or misassembled proteins. Chaperones, which are specialized proteins that assist in protein folding and assembly, play a key role in this process. Chaperones can recognize misfolded or unfolded proteins and either help them refold correctly or target them for degradation by proteases. In addition, cells have other quality control mechanisms, such as the unfolded protein response and the heat shock response, which are activated when cells are exposed to stress or when protein assembly is disrupted. These responses can help cells cope with misfolded or misassembled proteins and prevent them from accumulating and causing damage.

The role of signaling in the monitoring process

Signaling is involved in monitoring the assembly process of proteins, including RNR enzymes. The process of protein assembly is tightly regulated, and various signaling pathways are involved in ensuring that the correct proteins are produced, properly folded, and assembled into functional complexes. For example, in the case of RNR enzymes, the assembly process is regulated by several different proteins and signaling pathways. One important pathway involves the use of chaperones, which are specialized proteins that help to fold other proteins into their correct three-dimensional structure. Chaperones recognize misfolded or partially folded proteins and either assist in their folding or target them for degradation. Another important signaling pathway involves the use of ubiquitin, a small protein that is attached to target proteins to mark them for degradation by the proteasome. If errors occur during the assembly process, proteins may be misfolded or partially assembled, and they may be targeted for degradation by the proteasome via the ubiquitin pathway. The process of protein assembly is highly regulated and involves the coordinated action of multiple signaling pathways to ensure that the final product is properly folded, assembled, and functional.

Thioredoxin Reduces Ribonucleotide Reductase

The synthesis and maintenance of DNA, as well as the production of deoxyribonucleotides required for DNA synthesis, depend on the activities of RNR, Trx, and TrxR enzymes working together. The RNR enzyme provides the deoxyribonucleotides necessary for DNA synthesis, while the Trx and TrxR enzymes maintain the proper redox state of the RNR enzyme to ensure its proper function. These processes are interdependent and require coordinated activity of all three enzymes.

On a side note: A redox state refers to the oxidation-reduction state of a chemical system, which describes whether a chemical species has lost or gained electrons in a reaction. In a redox reaction, one reactant loses electrons (is oxidized) and another reactant gains electrons (is reduced). The overall process is called a redox reaction, which involves a transfer of electrons between molecules. The redox state of a system is important in many biochemical processes, including cellular respiration and photosynthesis. The redox state is important because it is involved in many biochemical processes, including energy production, biosynthesis, and signal transduction. In particular, many enzymes and proteins require a specific redox state to function properly. For example, the redox state of the electron transport chain in mitochondria is critical for ATP production, and the redox state of proteins involved in signaling pathways can affect gene expression and other cellular processes. Additionally, the redox state can play a role in oxidative stress and disease, as an imbalance in the redox state can lead to damage to cellular components and disruption of normal cellular function.

Thioredoxin plays a crucial role in the function of ribonucleotide reductase (RNR) enzymes.  Thioredoxin acts as a reducing agent, providing the electrons necessary for the reduction of the active site cysteine residues in RNR enzymes.

As a side note: In biochemistry, reducing refers to the process of donating electrons or hydrogen atoms to another molecule or substance, resulting in a decrease in its oxidation state. This can also be thought of as the process of removing oxygen from a molecule or adding hydrogen to it. Reduction reactions are essential for many biological processes, including cellular respiration, photosynthesis, and the synthesis of many important biomolecules such as amino acids, fatty acids, and nucleotides. Reduction reactions are typically carried out by enzymes that facilitate the transfer of electrons or hydrogen atoms between molecules. One example of a reducing agent in biochemistry is NADH (nicotinamide adenine dinucleotide), which donates electrons to other molecules during cellular respiration to produce ATP, the primary source of energy for the cell. Another example is glutathione, which is involved in reducing oxidative stress by donating electrons to reactive oxygen species (ROS) to neutralize them.

Specifically, thioredoxin reduces a disulfide bond in the RNR enzyme which is critical for its activity. This reduction allows the RNR enzyme to convert ribonucleotides to deoxyribonucleotides. In addition to its role as a reducing agent, thioredoxin also interacts with RNR enzymes to regulate their activity. For example, thioredoxin can bind to an allosteric site on RNR and activate the enzyme.

Thioredoxin NADPH Reductase (TrxR)

The redox function of Thioredoxin is critically dependent on the enzyme Thioredoxin NADPH Reductase (TrxR). Thioredoxin (Trx) and Thioredoxin NADPH Reductase (TrxR) are interdependent and work together to maintain cellular redox homeostasis. TrxR is an enzyme that catalyzes the transfer of electrons from NADPH to Trx, reducing Trx and maintaining it in its active reduced form. Trx, in turn, acts as a reducing agent for a wide range of cellular processes, including DNA synthesis, protein folding, and antioxidant defense. Without TrxR, Trx would remain in its oxidized form and be unable to carry out its essential functions. Conversely, without Trx, TrxR would be unable to carry out its function of transferring electrons to downstream targets, leading to a buildup of oxidized substrates. Trx and TrxR form an important redox couple in the cell, working together to maintain the cellular redox balance and enable a wide range of cellular processes. Trx and TrxR are found in all three domains of life - Bacteria, Archaea, and Eukarya - and share significant sequence and structural similarity across these domains.  These enzymes had to be fully developed for life to start.

Thioredoxin (Trx) and Thioredoxin NADPH Reductase (TrxR) are both proteins with specific structures that enable their biological function. Trx is a small protein that contains a conserved CXXC motif (where C is cysteine and X is any amino acid) that is essential for its redox activity.

In biology, the term "conserved" refers to the degree of similarity or identity between sequences or structures of biological molecules that are found in different organisms.

On a side note: When a sequence or structure is described as conserved, it means that it has remained relatively unchanged over evolutionary time and is therefore shared by many different organisms. This suggests that the sequence or structure is functionally important and has been conserved due to its essential role in biological processes. For example, the CXXC motif in Thioredoxin (Trx) is conserved across all domains of life, meaning that it has remained relatively unchanged and is present in Trx proteins from bacteria, archaea, and eukaryotes. This suggests that the CXXC motif is functionally important for Trx's redox activity and has been conserved due to its essential role in cellular processes. Similarly, the thioredoxin fold, which is the conserved three-dimensional structure adopted by Trx, is found in many other proteins that have diverse functions. This suggests that the thioredoxin fold is a versatile structural motif that has been conserved due to its functional importance.

Trx adopts a conserved three-dimensional fold called a thioredoxin fold, which consists of a central β-sheet flanked by several α-helices. The active site of Trx, where it interacts with other proteins and substrates, is located within a groove on the surface of the protein.

The active site of Thioredoxin (Trx) is a specific region on the surface of the protein where it interacts with other proteins and substrates, and where its redox activity occurs. It is located within a groove on the surface of the protein and is formed by the conserved CXXC motif. This motif contains two cysteine residues (Cys) that are separated by two other amino acids, and together they form a disulfide bond (S-S) that is essential for Trx's redox activity. The active site of Trx can interact with a variety of different substrates, including other proteins, enzymes, and DNA. Trx binds to these substrates through a combination of electrostatic, hydrophobic, and van der Waals interactions, as well as through specific recognition motifs or binding domains. In addition to its redox activity, the active site of Trx is also involved in protein-protein interactions and can interact with several different proteins and enzymes in the cell. These interactions can modulate the activity of Trx and its downstream targets, and play important roles in regulating cellular processes such as DNA synthesis, protein folding, and cell signaling.

The active-site surface in thioredoxin is designed to fit many proteins. Thioredoxin thus uses a chaperone-like mechanism of conformational changes to bind a diverse group of proteins and fast thiol-disulfide exchange chemistry in a hydrophobic environment to promote high rates of disulfide reduction.

TrxR, on the other hand, is a larger protein with a molecular weight of about 55-60 kDa. It is composed of two identical subunits that each contain a flavin adenine dinucleotide (FAD) cofactor, an NADPH binding site, and a redox-active disulfide bond. TrxR also contains a conserved C-terminal domain that interacts with Trx and transfers electrons from NADPH to Trx.

The crystal structures of Trx and TrxR have been extensively studied, and their structures are well-characterized. Understanding the structure of these enzymes is important for understanding their function and how they interact with other proteins and substrates in the cell.



1. Kinga Nyíri: Structural model of human dUTPase in complex with a novel proteinaceous inhibitor 12 March 2018
2. Jayachandran: Why deoxyribose for DNA and ribose for RNA? 2014
3. Gerald F. Joyce: The antiquity of RNA-based evolution 11 July 2002
4. Matthew Cobb: Life's Greatest Secret: The Race to Crack the Genetic Code page 178 7 julho 2015
5. Vinod Thakur: Why Nature Preferred DNA over RNA?  April 7, 2018
6. Marcos Eberlin: Foresight: How the Chemistry of Life Reveals Planning and Purpose  26 abril 2019
7. what-when-how:  In Depth Tutorials and Information Ribonucleotide Reductases (Molecular Biology)
8. Audrey A Burnim et.al.,: Comprehensive phylogenetic analysis of the ribonucleotide reductase family reveals an ancestral clade Sep 1, 2022
9. Daniel Lundin: The Origin and Evolution of Ribonucleotide Reduction 2015 Mar
10. Anders Hofer: DNA building blocks: keeping control of manufacture 03 Nov 2011
11. Soo-Cheul Yoo: Rice Virescent3 and Stripe1 Encoding the Large and Small Subunits of Ribonucleotide Reductase Are Required for Chloroplast Biogenesis during Early Leaf Development 1, May 2009
12. Pär Nordlund: Ribonucleotide reductases 2006
13. Reginald H. Garrett: Biochemistry 2016
14. Raleigh McElvery MIT News: Newly discovered enzyme “square dance” helps generate DNA building blocks March 30, 2020
15. Gyunghoon Kang: Structure of a trapped radical transfer pathway within a ribonucleotide reductase holocomplex 2020 Apr 24
16. Anne Trafton, MIT News: Chemists discover how a single enzyme maintains a cell’s pool of DNA building blocks. January 12, 2016
17. Evolution News: An "An "Exquisitely Designed" Enzyme that Maintains DNA Building Blocks January 16, 2016
18. Catherine L Drennan: Molecular basis for allosteric specificity regulation in class Ia ribonucleotide reductase from Escherichia coli JAN 12 2016
19. Edward J. Brignole: The prototypic class Ia ribonucleotide reductase from Escherichia coli: still surprising after all these years  2018 Apr 23
20. Eduard Torrents: Ribonucleotide reductases: essential enzymes for bacterial life 2014 Apr 28
21. Terry B. Ruskoski: The periodic table of ribonucleotide reductases OCTOBER 2021
22. Daniel Lundin: RNRdb, a curated database of the universal enzyme family ribonucleotide reductase, reveals a high level of misannotation in sequences deposited to Genbank 2009 Dec 8
23. Reginald H. Garrett: Biochemistry 4th ed. 2008 Pg.866
24. Anders Hofer: DNA building blocks: keeping control of manufacture 2012
25. Wikipedia: Ribonucleotide reductase
26. Alberts B et al., Molecular Biology of the Cell. 4th ed 2002.
27. Chabes, A., & Thelander, L. . Controlled protein degradation regulates ribonucleotide reductase activity in proliferating mammalian cells during the normal cell cycle and in response to DNA damage and replication blocks. 2000
28. Lundin, D., et.al.,  Ribonucleotide reduction—horizontal transfer of a required function spans all three domains. 2010
29. Lander, E. S., et.al,   Initial sequencing and analysis of the human genome. 2001
30. Johansson, R., et.al. Structures and mechanism of class I ribonucleotide reductase.  2021

The synthesis of Thymine Nucleotides





RNA contains the base uracil, which differs from thymine, the equivalent base in DNA, by the absence of a –CH3  ( methyl group ) Spontaneous deamination of cytosine to uracil in DNA occurs at a rate of about 100 bases per cell per day, and one of the most common methods of damage. ( see below ) Had DNA not switched from uracil to thymine, the deamination damage to cytosine would be essentially impossible to detect. But since thymine is used by DNA, uracil can be correctly recognized as damaged and repaired back to cytosine with thymine as template.



The thymine-uracil exchange constitutes one of the major chemical differences between DNA and RNA. Although these two bases form the same Watson-Crick base pairs with adenine and are equivalent for both information storage and transmission, uracil incorporation in DNA is usually a mistake that needs to be excised. There are two ways for uracil to appear in DNA: thymine replacement and cytosine deamination. 20 Most DNA polymerases readily incorporate dUMP as well as dTMP depending solely on the availability of the d(U/T)TP building block nucleotides. Cytosine deamination results in mutagenic U:G mismatches that must be excised. The repair system, however, also excises U from U:A “normal” pairs. It is therefore crucial to limit thymine-replacing uracils.

De novo biosynthesis of thymine is an intricate and energetically expensive process that requires dUMP as the starting material and a complex array of two enzymes and cofactors. It is therefore straightforward to ask: is there any specific reason that justifies this costly and seemingly equivalent replacement of uracil by thymine in DNA? It is generally accepted that negative discrimination against uracil in DNA is caused by the chemical instability of cytosine. Deamination of cytosine, a rather frequent process that readily occurs under physiological circumstances, gives rise to uracil .   Unless corrected, this mutagenic transition will result in a C:G into U(T): A base-pair change, that is, a stable point mutation. To deal with this problem, a highly efficient repair process (uracil-excision repair)  starts with uracil–DNA glycosylase (UDG). The importance of this repair process is well-reflected in two observations. One, cytosine deamination is one of the most frequent spontaneous mutations in DNA.Two, UDG activity resides in at least four families of enzymes: redundancy may be required for specific circumstances. 21

During nucleotide synthesis,  ribonucleotides that form RNA, are transformed into deoxyribonucleotides that form DNA. Before being incorporated into the chromosomes, another essential modification takes place. Uracil bases in RNA are transformed into thymine bases in DNA. There is a life essential requirement, why. Cytosine, the second of the two pyrimidine bases used in DNA, do deaminate over time into uracil bases. If uracil would remain, and not be replaced by thymine in DNA,  it would mix with the cytosine which deaminated spontaneously into uracil -  occurring on average, 100 times per day in the cell. Deamination of cytosine into deoxyuridine (a common spontaneous chemical reaction) can lead to incorporation of numerous mutations in the chromosome during replication with disastrous outcomes. If there wasn't a mechanism to remove the deaminated nucleotides (dUMP's), then, gradually over time, all of the Cytosine-Adenine base pairings would become a Uracil-Adenine base pairing.

If uracil would be transformed from RNA to DNA, transforming it into deoxy uracil, and used in DNA like it is in RNA, and not replaced with thymine, then it would keep being recognized as deaminated cytidine by the repair machinery, and removed as well,  and the DNA would be basically coated in uracil DNA glycosylases repair enzymes removing the deaminated base pairs, and the legitimate ones. So, instead, DNA uses thymidine, which is distinguishable biochemically from uracil by its extra methyl group. This way the cell gains the essential ability to remove the uracils that are a result of deamination using uracil DNA glycosylase enzymes, preventing mass mutation in its genome without removing the thymine base that it actually needs to be there.

The buildup of these “illegitimate” uracils could be catastrophic for the organism - at the very least, copying fidelity of DNA would be detrimentally affected. Thus, cells have repair systems in place to remove these “illegitimate” uracils. But if uracil were already present in DNA, paired to adenine, the repair system would be forced to somehow differentiate between “illegitimate” and “legitimate” uracils. An easy solution to this problem? Add a methyl group to all of the “legitimate” uracils, allowing the repair system to easily tell between the two. This usage of methylated uracil, or thymine, in DNA allowed for the long-term storage of crucial genetic information. 6

In a DNA organism, deoxyuridine can naturally be distinguished from thymidine and be repaired to cytosine. This process cannot take place in the case of RNA deamination, highlighting the great advantage of the invention of thymidine.

But why would prebiotic molecules without distant goals nor purpose to produce a stable information storage medium, DNA, promote this base exchange, and produce error check and repair mechanisms to keep the information intact and promote high-fidelity replication and maintain the mutation levels low?

Only one extra synthetic step in nucleotide biosynthesis is required to achieve the exchange of uracil to thymine, but the machinery to do the job is enormously complex.

After further phosphorylation, that is, adding two phosphate groups to the deoxynucleotide monophosphates, they become deoxynucleotide triphosphates dGTP, dATP, dCTP, and dTTP and can be used as the building blocks to construct DNA.

The deoxynucleotide triphosphates (dNTPs) are the building blocks for DNA replication (they lose two of the phosphate groups in the process of incorporation and polymerization). 7  Phosphorylation status of nucleotides is regulated by NDP kinases and NMP kinases that use ATP pool as their cross-phosphorylation source.




Thymidine is a deoxyuridine with a methyl group at the C5 position of the uracil base. This subtle difference plays a critical role in the superior fidelity of DNA-based replication over its RNA counterpart.
Methylation protects the DNA. Beside using thymine instead of uracil, most organisms also use various enzymes to modify DNA after it has been synthesized. Two such enzymes, dam and dcm methylate adenines and cytosines, respectively, along the entire DNA strand. This methylation makes the DNA unrecognizable to many Nucleases (enzymes which break down DNA and RNA), so that it cannot be easily attacked by invaders, like viruses or certain bacteria. Obviously, methylating the nucleotides before they are incorporated ensures that the entire strand of DNA is protected.

Thymine also protects the DNA in another way. If you look at the components of nucleic acids, phosphates, sugars, and bases, you see that they are all very hydrophilic (water soluble). Obviously, adding a hydrophobic (water insoluble) methyl group to part of the DNA is going to change the characteristics of the molecule. The major effect is that the methyl group will be repelled by the rest of the DNA, moving it to a fixed position in the major groove of the helix. This solves an important problem with uracil - though it prefers adenine, uracil can base-pair with almost any other base, including itself, depending on how it situates itself in the helix. By tacking it down to a single conformation, the methyl group restricts uracil (thymine) to pairing only with adenine. This greatly improves the efficiency of DNA replication, by reducing the rate of mismatches, and thus mutations.

To sum up: the replacement of thymine for uracil in DNA protects the DNA from attack and maintains the fidelity of DNA replication.

RNA contains the base uracil, which differs from thymine, the equivalent base in DNA, by the absence of a –CH3  ( methyl group ) Spontaneous deamination of cytosine to uracil in DNA occurs at a rate of about 100 bases per cell per day, and one of the most common methods of damage. Had DNA not switched from uracil to thymine, the deamination damage to cytosine would be essentially impossible to detect. But since thymine is used by DNA, uracil can be correctly recognized as damaged and repaired back to cytosine with thymine as a template. The addition of the methyl group to thymine provides a way for the DNA repair machinery to distinguish between normal, methylated thymine and potentially mutagenic uracil that may have been incorporated into the DNA through various mechanisms. This helps to prevent errors in DNA replication and maintains the stability of the genome.

Cytosine deamination is a spontaneous chemical reaction that can occur in DNA and results in the conversion of cytosine (C) to uracil (U). Uracil is normally found in RNA but not in DNA. When uracil is present in DNA, it can form a base pair with guanine (G), creating a U:G mismatch that can cause mutations if not corrected. To repair such mismatches, cells have a specialized DNA repair pathway called base excision repair (BER), which removes the uracil base and replaces it with a cytosine base. However, BER is not specific to U:G mismatches and can also remove uracil from normal U:A base pairs. This is because BER recognizes uracil as an abnormal base that should not be present in DNA, regardless of whether it is paired with guanine or adenine.

Therefore, it is crucial to limit the occurrence of thymine-replacing uracils in DNA to prevent the unnecessary removal of uracil from normal U:A base pairs during BER. One way this is achieved is through the action of DNA methyltransferases, which add a methyl group to the C5 position of cytosine to form 5-methylcytosine (5mC). This modification makes cytosine less susceptible to deamination and reduces the occurrence of U:G mismatches. Another way to limit thymine-replacing uracils is through the action of DNA glycosylases, which specifically recognize and remove uracil from U:G mismatches while leaving normal U:A base pairs intact. This is possible because the DNA glycosylase enzymes can distinguish the difference between a normal U:A base pair and a U:G mismatch based on the local DNA structure and other factors.

The problem of preventing thymine-replacing uracils in DNA and recognizing U:G mismatches had to be fully solved at the time when life began,  and this solution had to be implemented instantly. De novo biosynthesis of thymine is a complex and energetically expensive process that requires the starting material, dUMP (2'-deoxyuridine 5'-monophosphate), and a series of enzymes and cofactors to convert it into thymine. This process occurs in the cytoplasm of most organisms, including bacteria, archaea, and eukaryotes.

Deoxyuridine 5′-triphosphate nucleotidohydrolase (dUTPase) 

The function of Deoxyuridine 5′-triphosphate nucleotidohydrolase (dUTPase) is to maintain the fidelity of DNA replication by preventing the incorporation of uracil into DNA in place of thymine. dUTP is a nucleotide that is structurally similar to dTTP (deoxythymidine triphosphate), which is the nucleotide that is normally used to pair with adenine during DNA replication. However, if dUTP is mistakenly incorporated into DNA in place of dTTP, it can lead to the formation of uracil-DNA, which can be mutagenic and result in DNA damage. dUTPase catalyzes the hydrolysis of dUTP to dUMP (deoxyuridine monophosphate) and inorganic pyrophosphate. By reducing the levels of dUTP in the cell, dUTPase helps to prevent its incorporation into DNA and reduce the potential for mutagenesis. dUTPase plays a critical role in maintaining the fidelity of DNA replication by preventing the incorporation of uracil into DNA and promoting the use of thymine instead.

The de novo biosynthesis of thymine nucleotides




Biosynthesis of thymidylate (dTMP)
The pathways are shown beginning with the reaction catalyzed by ribonucleotide reductase. Figure above gives details of the thymidylate synthase reaction.

The first step in this pathway is the conversion of dUMP to dTMP (2'-deoxythymidine 5'-monophosphate) by the enzyme thymidylate synthase. This reaction requires a cofactor called 5,10-methylenetetrahydrofolate (5,10-MTHF), which donates a methyl group to dUMP to form dTMP. 5,10-MTHF is itself synthesized from folic acid through a series of reactions that involve several enzymes and cofactors.

The second step in the pathway is the phosphorylation of dTMP to form dTDP (2'-deoxythymidine 5'-diphosphate) by the enzyme thymidylate kinase. This reaction requires the input of ATP (adenosine triphosphate) as a source of phosphate groups. The final step in the pathway is the conversion of dTDP to dTTP (2'-deoxythymidine 5'-triphosphate) by the enzyme nucleoside diphosphate kinase, which uses ATP as a source of phosphate groups. dTTP is the final product of the pathway and is used as a building block for DNA synthesis. Overall, the de novo biosynthesis of thymine is a complex process that requires the input of several enzymes and cofactors, as well as significant amounts of energy in the form of ATP. This pathway is essential for the synthesis of DNA and the maintenance of genomic stability, and its dysregulation is associated with several diseases, including cancer and autoimmune disorders.

The de novo biosynthesis of thymine involves several enzymes, including:

1. Thymidylate synthase (TS): catalyzes the conversion of dUMP to dTMP. This enzyme requires the cofactor 5,10-methylenetetrahydrofolate (5,10-MTHF) as a source of methyl groups.
2. Dihydrofolate reductase (DHFR): catalyzes the conversion of dihydrofolate (DHF) to 5,10-MTHF, which is required as a cofactor for TS.
3. Thymidylate kinase (TMPK): phosphorylates dTMP to form dTDP, using ATP as a source of phosphate groups.
4. Ucleoside diphosphate kinase (NDPK): converts dTDP to dTTP, using ATP as a source of phosphate groups.

Furthermore, following Cofactors are also required: 

NADPH (nicotinamide adenine dinucleotide phosphate, reduced form)
ATP (adenosine triphosphate)
GTP (guanosine triphosphate)
5,10-methylenetetrahydrofolate (5,10-MTHF)

Thymine has an additional benefit for DNA beyond just protecting it from deamination. The addition of a hydrophobic methyl group to thymine changes the chemical characteristics of the molecule, making it water-insoluble and causing it to be repelled by the rest of the DNA. This moves the methyl group to a fixed position in the major groove of the DNA helix. This positioning of the methyl group has an important effect on the base-pairing properties of thymine. Uracil, which is a chemically similar base to thymine, can pair with almost any other base depending on its conformation in the helix. This means that during DNA replication, there is a higher risk of mismatches and mutations if uracil is present in the DNA. By tacking down thymine to a fixed conformation through the methyl group, it restricts thymine to base-pair only with adenine. This greatly improves the efficiency and accuracy of DNA replication by reducing the rate of mismatches and mutations. The replacement of uracil with thymine in DNA not only protects the DNA from deamination but also helps to maintain the fidelity of DNA replication. The addition of the hydrophobic methyl group to thymine restricts it to base-pair only with adenine, reducing the rate of mismatches and mutations during DNA replication.

Thymidylate synthase (TS)

Thymidylate synthase (TS) is an essential enzyme found in all living cells today.  It is involved in the de novo synthesis of DNA. It catalyzes the conversion of deoxyuridine monophosphate (dUMP) to deoxythymidine monophosphate (dTMP), which is a precursor for thymidine, one of the four nucleotides that make up DNA. Thus, TS plays a critical role in DNA replication and cell division. If a cell were to completely lack thymidylate synthase (TS) activity, it would not be able to produce deoxythymidine triphosphate (dTTP), which is an essential building block for DNA synthesis. As a result, the cell would not be able to replicate its DNA properly, and DNA synthesis would eventually come to a halt. This would lead to cell cycle arrest and eventually cell death, as the cell would not be able to maintain its genetic material or carry out essential cellular functions that require DNA replication.

The smallest version of thymidylate synthase (TS) is found in some bacterial species and consists of only a single polypeptide chain. These bacterial TS enzymes are known as ThyA enzymes and are typically around 180 to 190 amino acids in length, with a molecular weight of approximately 20 kDa. Unlike the more complex TS enzymes found in higher organisms, ThyA enzymes do not require a separate folate cofactor to function, but instead rely on the amino acid methionine as a source of one-carbon units. The smaller size of bacterial ThyA enzymes is thought to reflect the simpler metabolic needs of these organisms compared to more complex eukaryotic organisms.

Thymidylate synthase (TS) is a dimeric enzyme consisting of two identical subunits, each with a molecular weight of approximately 30 kDa. The amino acid sequence of TS varies between species, but in humans, the protein is composed of 313 amino acids. TS has a complex secondary structure consisting of eight alpha helices and eight beta strands arranged in a barrel-like structure known as a TIM barrel. The TIM barrel is a common feature of many enzymes involved in metabolic pathways, and it provides a stable framework for the active site of the enzyme. The active site of TS is located at the C-terminal end of the protein and is composed of several amino acid residues that are critical for enzyme function. These residues include a conserved cysteine residue that is involved in the formation of a covalent bond with the substrate, as well as several other amino acids that help to orient the substrate and catalyze the reaction. In addition to the TIM barrel domain, TS also contains a unique insert domain that is involved in binding to the cofactor, 5,10-methylenetetrahydrofolate (MTHF). The insert domain is located at the N-terminal end of the protein and is composed of approximately 25 amino acids that form a loop structure. This loop interacts with MTHF and helps to position it correctly in the active site of the enzyme. Overall, the structure of TS is complex and involves several distinct domains that are critical for enzyme function. The TIM barrel provides a stable framework for the active site, while the insert domain helps to bind the cofactor and position it correctly for catalysis.

Mechanism description

The mechanism of thymidylate synthase (TS) involves the transfer of a methyl group from 5,10-methylenetetrahydrofolate (CH2-THF) to deoxyuridine monophosphate (dUMP), which results in the formation of deoxythymidine monophosphate (dTMP), a key precursor for the synthesis of DNA. The reaction occurs in two steps, as follows:

Step 1: TS catalyzes the transfer of a hydride ion from dUMP to CH2-THF, generating dihydrofolate (DHF) and a covalent TS-dUMP intermediate.
Step 2: The covalent intermediate is then attacked by a nucleophilic thiol group on the enzyme, which results in the transfer of the methyl group from CH2-THF to dUMP, generating dTMP and releasing the enzyme.

In addition to its catalytic activity, TS is also involved in the regulation of DNA synthesis through a feedback mechanism that involves binding of the end product dTMP to the enzyme. Binding of dTMP to TS inhibits the enzyme's activity, thereby reducing the production of dTMP and slowing down DNA synthesis. This feedback mechanism helps to ensure that the cell maintains appropriate levels of dTMP for DNA synthesis and avoids overproduction of this essential precursor.

5,10-methylenetetrahydrofolate (CH2-THF) is a derivative of folic acid, which is a vitamin that is essential for many cellular processes, including DNA synthesis, repair, and methylation. In cells, folic acid is converted to tetrahydrofolate (THF), which is a key coenzyme that carries one-carbon units involved in a variety of metabolic reactions, including nucleotide synthesis.

The formation of CH2-THF involves the activity of the enzyme serine hydroxymethyltransferase, which transfers a methyl group from serine to THF, generating CH2-THF and glycine. CH2-THF can then be used as a source of one-carbon units in a variety of reactions, including the synthesis of thymidylate by TS.

Folic acid is obtained from the diet and is commonly found in leafy green vegetables, legumes, and fortified cereals. However, some organisms, such as bacteria and plants, are able to synthesize folic acid de novo, whereas others, such as humans, must obtain it from dietary sources.

Thymidylate synthase (TS) is an enzyme that requires the coenzyme tetrahydrofolate (THF), which is derived from folic acid, in order to function. Specifically, TS catalyzes the conversion of deoxyuridine monophosphate (dUMP) to deoxythymidine monophosphate (dTMP), a critical step in DNA synthesis. THF is necessary for TS to transfer a methyl group to dUMP, forming dTMP. Without THF, TS cannot function, and the cell's ability to synthesize DNA would be impaired.

Folic acid

Folic acid (also known as folate or vitamin B9) is essential for life as it plays a critical role in many cellular processes, including DNA synthesis, repair, and methylation. Folic acid is a key co-factor in the transfer of one-carbon units, which are required for the biosynthesis of nucleotides, amino acids, and other important molecules. 

Synthesis of folic acid

Folic acid is synthesized in cells through a complex pathway involving several enzymes and co-factors. The pathway is highly conserved across different organisms, from bacteria to mammals. Here is a brief overview of the folic acid synthesis pathway in bacteria:

1. The first step in the pathway involves the synthesis of dihydropteroate from para-aminobenzoic acid (PABA) by the enzyme dihydropteroate synthase.
2. Dihydropteroate is then converted to dihydrofolate (DHF) by the enzyme dihydrofolate reductase.
3. DHF is then converted to tetrahydrofolate (THF) by the enzyme dihydrofolate reductase.
4. THF is then modified by the addition of various one-carbon units, including methyl, formyl, and methylene groups, which are derived from serine, histidine, and glycine, respectively. These modifications are catalyzed by a series of enzymes, including serine hydroxymethyltransferase, methylenetetrahydrofolate reductase, and formyltetrahydrofolate synthase.
5. The final step in the pathway involves the synthesis of folic acid from THF by the addition of a glutamate residue, which is catalyzed by the enzyme dihydrofolate synthase.

The biosynthesis pathway of folic acid involves several enzymes, co-factors, and substrates, and the number of enzymes involved varies among different organisms. In bacteria, the pathway is relatively simple and involves approximately 7-8 enzymes, whereas in humans and other mammals, the pathway is more complex and involves at least 15 enzymes. The enzymes involved in the pathway catalyze a variety of reactions, including condensations, reductions, methylations, and formylations, and are essential for the production of the various forms of folate needed for many cellular processes, including DNA synthesis, repair, and methylation. All the enzymes involved in the biosynthesis of folic acid are essential for life, as they are involved in critical metabolic pathways that are necessary for normal cellular function. Deficiencies in any of these enzymes can lead to impaired folic acid synthesis, which can result in a range of health problems.  Some of the enzymes involved in folic acid biosyntheses, such as dihydropteroate synthase and dihydrofolate reductase, are ancient and have been found in many different organisms, suggesting that they may have been present in the last universal common ancestor (LUCA).

The smallest pathway for the biosynthesis of folic acid is found in some bacteria and consists of approximately 7-8 enzymes. The sizes of the enzymes in this pathway vary, but some of the smallest enzymes are:

1. GTP cyclohydrolase I (GCHI) - This enzyme catalyzes the conversion of GTP to dihydroneopterin triphosphate, which is the first step in the biosynthesis of folate. GCHI is a relatively small enzyme consisting of approximately 240-250 amino acids, depending on the organism.
2. 6-hydroxymethyl-7,8-dihydropterin pyrophosphokinase (HPPK) - This enzyme catalyzes the conversion of dihydroneopterin triphosphate to 6-hydroxymethyl-7,8-dihydropterin pyrophosphate. HPPK is a relatively small enzyme consisting of approximately 200-250 amino acids, depending on the organism.
3. Dihydropteroate synthase (DHPS) - This enzyme catalyzes the condensation of 6-hydroxymethyl-7,8-dihydropterin pyrophosphate with p-aminobenzoic acid (PABA) to form dihydropteroate, which is a precursor to folate. DHPS is a relatively small enzyme consisting of approximately 250-300 amino acids, depending on the organism.
4. Dihydrofolate synthase (DHFS) - This enzyme catalyzes the reduction of dihydropteroate to dihydrofolate, which is the immediate precursor to tetrahydrofolate. DHFS is a relatively small enzyme consisting of approximately 200-250 amino acids, depending on the organism.
5. Dihydrofolate reductase (DHFR) - This enzyme catalyzes the reduction of dihydrofolate to tetrahydrofolate, which is the active form of folate used in cellular metabolism. DHFR is a relatively small enzyme consisting of approximately 150-200 amino acids, depending on the organism.

Note that the exact sizes of these enzymes can vary depending on the organism and the specific isoform of the enzyme. Based on the estimated sizes of the enzymes outlined in the smallest pathway for the biosynthesis of folic acid, the total number of amino acids would be approximately 1040-1200, depending on the organism and the specific isoforms of the enzymes involved. the enzymes in the biosynthesis pathway for folic acid must be arranged in the correct order and work together as a production line to synthesize folic acid. Each enzyme performs a specific reaction that converts one intermediate molecule to the next in the pathway, and the products of one enzyme become the substrates for the next enzyme in the sequence. Any disruption in the order or activity of the enzymes in the pathway can lead to a deficiency in folate synthesis and affect cellular processes that require folate. The probability of the correct functional enzymes arising by chance is already low, and the probability of lining them up in the correct order is even lower. The correct order and arrangement of enzymes are essential for the efficient and accurate biosynthesis of folic acid. The probability of random processes producing the right sequence of functional enzymes for the biosynthesis of folic acid is extremely low, which makes it difficult to explain the origin of this pathway solely through chance events. Given that the folic acid biosynthesis pathway consists of multiple enzymes that must be arranged in a specific order to produce the end product, the probability of this pathway arising through a series of chance events is even lower. In other words, the odds of a chance event producing a metabolic production line like the folic acid biosynthesis pathway are so low that it is considered highly unlikely, if not impossible, based on current scientific understanding.

There is an interdependence between thymidylate synthase (TS) and folic acid. TS requires folic acid-derived tetrahydrofolate (THF) as a coenzyme to function, and without THF, TS cannot carry out its enzymatic activity. Conversely, folic acid relies on the activity of TS to be converted into its active coenzyme form, THF. Therefore, the functions of TS and folic acid are interdependent.

In addition to tetrahydrofolate (THF), thymidylate synthase (TS) also requires the cofactor N5,N10-methylene-tetrahydrofolate (CH2-THF) and the reducing agent NADPH to function. CH2-THF serves as the source of the methyl group that is transferred to deoxyuridine monophosphate (dUMP) to form deoxythymidine monophosphate (dTMP). NADPH donates the electrons required to reduce CH2-THF to form the active methyl donor, CH3-THF. Without CH2-THF or NADPH, TS cannot function.

N5,N10-methylene-tetrahydrofolate (CH2-THF)

N5,N10-methylene-tetrahydrofolate (CH2-THF) is a coenzyme that serves as the source of the methyl group in the enzymatic reaction catalyzed by thymidylate synthase (TS). The methyl group is transferred from CH2-THF to deoxyuridine monophosphate (dUMP) to form deoxythymidine monophosphate (dTMP), which is an essential precursor for DNA synthesis. CH2-THF is a derivative of tetrahydrofolate (THF), which is synthesized from folic acid. The methylene group in CH2-THF is derived from serine through the action of the enzyme serine hydroxymethyltransferase. In summary, CH2-THF plays a critical role in the biosynthesis of DNA, and its production depends on the availability of folic acid and serine.

Synthesis of N5,N10-methylene-tetrahydrofolate (CH2-THF)

N5,N10-methylene-tetrahydrofolate (CH2-THF) is synthesized through a series of enzymatic reactions that involve the conversion of tetrahydrofolate (THF) to N5,N10-methylenetetrahydrofolate (CH2-THF). The conversion is catalyzed by the enzyme methylenetetrahydrofolate reductase (MTHFR), which reduces N5,N10-methylenetetrahydrofolate (CH2-THF) from N5,N10-methylenetetrahydrofolate (CH2-THF) by transferring electrons from NADPH. The reaction requires the presence of vitamin B12 as a cofactor.

The biosynthesis of N5,N10-methylene-tetrahydrofolate (CH2-THF) starts with the conversion of folic acid to dihydrofolate (DHF) by the enzyme dihydrofolate reductase (DHFR). DHF is then reduced to THF by DHFR or another enzyme, depending on the cell type. THF is then converted to N5,N10-methylenetetrahydrofolate (CH2-THF) by MTHFR.

It's important to note that the production of CH2-THF is dependent on the availability of folic acid, which is obtained through dietary sources in humans and other animals. In summary, CH2-THF is synthesized from THF through the action of MTHFR, and its production is dependent on the availability of folic acid and vitamin B12.

The smallest pathway to obtain N5,N10-methylene-tetrahydrofolate (CH2-THF) involves two enzymes:

Dihydrofolate reductase (DHFR) - This enzyme reduces dihydrofolate (DHF) to tetrahydrofolate (THF). The smallest known DHFR enzyme is found in the bacterium Mycoplasma genitalium, which has a size of approximately 17 kiloDaltons (kDa) and a length of approximately 159 amino acids. It is an enzyme that plays a crucial role in folate metabolism by catalyzing the reduction of dihydrofolate (DHF) to tetrahydrofolate (THF), which is essential for DNA synthesis, repair, and methylation. The reduction reaction is a two-step process that involves the transfer of two electrons and one proton from NADPH to DHF, resulting in the formation of THF and NADP+. THF serves as a coenzyme for various enzymes involved in one-carbon metabolism, which is critical for the synthesis of nucleotides, amino acids, and other biomolecules. DHFR is essential for cell growth and division It requires a cofactor called NADPH (nicotinamide adenine dinucleotide phosphate) to work. NADPH donates electrons to DHFR during the reduction of dihydrofolate (DHF) to tetrahydrofolate (THF). Without NADPH, DHFR cannot perform its enzymatic activity.

Methylenetetrahydrofolate reductase (MTHFR) - This enzyme converts THF to N5,N10-methylene-tetrahydrofolate (CH2-THF). The smallest known MTHFR enzyme is found in the bacterium Escherichia coli, which has a size of approximately 26 kDa and a length of approximately 235 amino acids. Methylenetetrahydrofolate reductase (MTHFR) is an enzyme that plays a critical role in folate metabolism by catalyzing the conversion of 5,10-methylenetetrahydrofolate (CH2-THF) to 5-methyltetrahydrofolate (5-MTHF), which is a coenzyme that serves as a methyl donor in various cellular processes, including DNA methylation and neurotransmitter synthesis. The reaction involves the transfer of a methyl group from CH2-THF to homocysteine, resulting in the formation of 5-MTHF and methionine. Methionine is further converted to S-adenosylmethionine (SAM), which is a universal methyl donor that participates in numerous cellular methylation reactions. MTHFR is also involved in the remethylation of homocysteine to methionine, which is critical for the maintenance of normal homocysteine levels in the body. Methylenetetrahydrofolate reductase (MTHFR) is an enzyme that depends on several cofactors to operate. It requires flavin adenine dinucleotide (FAD) as a prosthetic group and uses NADPH as a reducing agent. Additionally, it depends on riboflavin (vitamin B2) and folate (vitamin B9) for its activity. Folate serves as a substrate for MTHFR and is converted to 5-methyltetrahydrofolate, which is necessary for the conversion of homocysteine to methionine. Therefore, adequate intake of vitamins B2 and B9 is important for the proper functioning of MTHFR

Riboflavin (vitamin B2)

Some bacteria, fungi, and plants can synthesize riboflavin. In bacteria, the synthesis of riboflavin involves a pathway of seven enzymes, while in plants and fungi, it involves a different pathway of six enzymes. The exact mechanisms of riboflavin synthesis vary depending on the organism, but they all involve a series of chemical reactions that convert precursor molecules into riboflavin.

Here is an outline of the enzymes involved in the riboflavin biosynthesis pathway, along with their sizes and amino acid lengths:

1. GTP cyclohydrolase II: This enzyme is a bifunctional enzyme that catalyzes the conversion of GTP to 2,5-diamino-6-ribosylamino-4(3H)-pyrimidinone 5'-phosphate (DRAPP), which is a precursor of riboflavin. Its size and amino acid length vary depending on the organism, but in bacteria, it is typically around 50 kDa and 450-500 amino acids in length.
2. Riboflavin synthase: This enzyme catalyzes the conversion of DRAPP to riboflavin. Its size and amino acid length also vary depending on the organism, but in bacteria, it is typically around 25 kDa and 230-260 amino acids in length.
3. Lumazine synthase: This enzyme catalyzes the formation of 6,7-dimethyl-8-ribityllumazine, which is an intermediate in the riboflavin biosynthesis pathway. Its size and amino acid length also vary depending on the organism, but in bacteria, it is typically around 45 kDa and 380-420 amino acids in length.
4. 6,7-dimethyl-8-ribityllumazine synthase: This enzyme catalyzes the conversion of 6,7-dimethyl-8-ribityllumazine to 7,8-dimethyl-8-ribityllumazine, which is another intermediate in the pathway. Its size and amino acid length also vary depending on the organism, but in bacteria, it is typically around 30 kDa and 270-300 amino acids in length.
5. 7,8-dimethyl-8-ribityllumazine phosphate synthase: This enzyme catalyzes the conversion of 7,8-dimethyl-8-ribityllumazine to 7,8-dimethyl-8-ribityllumazine 5'-phosphate. Its size and amino acid length also vary depending on the organism, but in bacteria, it is typically around 30 kDa and 270-300 amino acids in length.
6. Riboflavin kinase: This enzyme catalyzes the conversion of riboflavin to riboflavin 5'-phosphate, which is the biologically active form of the vitamin. Its size and amino acid length also vary depending on the organism, but in bacteria, it is typically around 25 kDa and 230-260 amino acids in length.

The total number of amino acids required for the entire pathway of riboflavin biosynthesis varies depending on the organism and the specific enzymes involved. However, a rough estimate, adding up the amino acids from each enzyme, the total comes to approximately 1,384 amino acids.  The pathway for synthesizing Riboflavin (vitamin B2) is essential for life, as the vitamin is necessary for various cellular processes, including energy production, growth, and development. Its origin is therefore related to the origin of life. 

In summary, the smallest pathway to obtain N5,N10-methylene-tetrahydrofolate (CH2-THF) involves two enzymes: DHFR and MTHFR, with the smallest known DHFR enzyme having approximately 159 amino acids, and the smallest known MTHFR enzyme having approximately 235 amino acids.

Dihydrofolate reductase (DHFR)

Dihydrofolate reductase (DHFR) is an enzyme that plays an important role in the metabolism of folate, a B vitamin that is essential for the synthesis of nucleic acids and amino acids. DHFR catalyzes the conversion of dihydrofolate (DHF) to tetrahydrofolate (THF), which is required for the synthesis of nucleic acids, and is also a co-factor for various metabolic reactions. DHFR is found in many organisms, including bacteria, fungi, plants, and animals, and is highly conserved across species.  DHFR is an important enzyme involved in the metabolism of folate, which is essential for the synthesis of nucleic acids and amino acids, and is a key target for certain drugs used in the treatment of cancer and infectious diseases.

The smallest version of dihydrofolate reductase (DHFR) is a bacterial enzyme known as E. coli DHFR. It is widely used as a model system for studying enzyme structure and function, and is the most extensively characterized DHFR enzyme. The size of E. coli DHFR is approximately 18 kDa, and it consists of 159 amino acids. It is a monomeric enzyme, meaning that it is composed of a single polypeptide chain. Despite its small size, E. coli DHFR is highly conserved across species and shares significant sequence similarity with DHFR enzymes from other organisms. If a cell were to lack the dihydrofolate reductase (DHFR) enzyme, it would not be able to convert dihydrofolate (DHF) to tetrahydrofolate (THF), which is essential for the synthesis of nucleic acids and amino acids. Without THF, the cell would not be able to produce nucleotides, which are the building blocks of DNA and RNA, and would not be able to synthesize certain amino acids, which are essential for protein synthesis. This would lead to impaired cell growth and division, as well as a range of other cellular functions that require nucleotides and amino acids. Therefore, the absence of DHFR can have a significant impact on cellular function and survival, and can ultimately lead to cell death.

Mechanism description

The mechanism of dihydrofolate reductase (DHFR) involves the reduction of dihydrofolate (DHF) to tetrahydrofolate (THF) using NADPH as a cofactor.  The mechanism of DHFR involves the transfer of a hydride ion (H-) from NADPH to DHF, which results in the reduction of DHF to THF. This reaction is catalyzed by a conserved set of amino acid residues within the active site of DHFR, which includes a highly reactive cysteine residue and a tyrosine residue that acts as a proton shuttle. The proton shuttle in dihydrofolate reductase (DHFR) is a conserved tyrosine residue that plays a critical role in the catalytic mechanism of the enzyme. The proton shuttle is responsible for the transfer of a proton from the N5 position of DHF to the C6 position of DHF during the reduction reaction, which is necessary for the formation of the reactive intermediate that leads to the production of tetrahydrofolate (THF).

The proton shuttle works in conjunction with a highly reactive cysteine residue within the active site of DHFR. The cysteine residue acts as a nucleophile, attacking the C6 position of DHF and forming a covalent bond with the substrate.

The meaning of "attacking" in biochemistry:  In biochemistry, "attacking" typically refers to the process of a nucleophile, such as the cysteine residue in dihydrofolate reductase (DHFR), forming a covalent bond with an electrophile, such as the C6 position of DHF.

What is a nucleophile? A nucleophile is a chemical species, such as an ion or a molecule, that has a tendency to donate a pair of electrons to an electrophile in order to form a chemical bond. Nucleophiles are characterized by the presence of a lone pair of electrons or a negative charge, which makes them attracted to positively charged or electron-deficient atoms or molecules.

What is a lone pair of electrons or a negative charge ?  An electron pair is a pair of electrons that are associated with an atom or molecule and are not involved in bonding with other atoms or molecules. In some cases, an atom or molecule may have one or more lone pairs of electrons, which are not involved in chemical bonding and are typically found in the outermost (valence) shell of the atom or molecule.

In the case of nucleophiles, a lone pair of electrons or a negative charge is important because it gives the molecule or ion a partial negative charge that makes it more attractive to positively charged or electron-deficient atoms or molecules. When a nucleophile encounters an electrophile (an atom or molecule that is electron-deficient), the nucleophile is able to donate its lone pair of electrons to form a new chemical bond with the electrophile.

For example, in the case of the cysteine residue in DHFR, the sulfur atom has a lone pair of electrons that can be used to attack the electrophilic carbon atom at the C6 position of DHF, leading to the formation of a new covalent bond between the cysteine residue and the substrate. The nucleophilic attack by the cysteine residue is an important step in the catalytic mechanism of DHFR, and is essential for the reduction of DHF to tetrahydrofolate (THF).

In biochemistry, nucleophiles play a critical role in a wide range of reactions, including enzyme-catalyzed reactions such as the one catalyzed by dihydrofolate reductase (DHFR). In DHFR, a cysteine residue acts as a nucleophile by donating a pair of electrons to the C6 position of dihydrofolate (DHF), which forms a covalent bond with the substrate and leads to the formation of a highly reactive intermediate.

Other examples of nucleophiles in biochemistry include the hydroxyl group in serine, threonine, and tyrosine residues, which play important roles in enzyme catalysis and protein function. The carboxylate group in aspartic acid and glutamic acid residues can also act as a nucleophile in certain enzyme-catalyzed reactions. Nucleophiles are also important in many other areas of chemistry, including organic synthesis and materials science.


In the context of DHFR, the cysteine residue acts as a nucleophile by donating a lone pair of electrons to the C6 position of DHF, which is an electrophilic carbon atom. This results in the formation of a covalent bond between the cysteine residue and the substrate, and leads to the formation of a highly reactive intermediate that is capable of accepting a hydride ion from NADPH.

The attack of the cysteine residue on the C6 position of DHF is an essential step in the catalytic mechanism of DHFR, and is necessary for the reduction of DHF to tetrahydrofolate (THF). The mechanism of DHFR is highly conserved across species, and is critical for the synthesis of nucleotides and certain amino acids.

This results in the formation of a highly reactive intermediate that is capable of accepting a hydride ion from NADPH. However, in order for the reaction to proceed, a proton must be transferred from the N5 position of DHF to the C6 position. This is where the proton shuttle comes into play. The conserved tyrosine residue within the active site of DHFR acts as a proton shuttle, transferring a proton from the N5 position of DHF to the C6 position, thereby facilitating the transfer of the hydride ion from NADPH to the substrate. This proton transfer is critical for the formation of the reactive intermediate, and is an essential step in the catalytic mechanism of DHFR.

The importance of the proton shuttle in the catalytic mechanism of DHFR is highlighted by the fact that mutations in the tyrosine residue can lead to a loss of enzyme activity, and can result in a range of health problems. 

The reaction proceeds through a series of steps involving the formation of a ternary complex between DHF, NADPH, and the enzyme. 

What is a ternary complex? A ternary complex is a complex formed between three molecules, typically an enzyme, a substrate, and a cofactor or inhibitor. In biochemistry, ternary complexes are important in many enzyme-catalyzed reactions, where they play a critical role in regulating the activity of enzymes and controlling the flow of metabolic pathways. In the context of enzyme catalysis, a ternary complex typically refers to the complex formed between an enzyme, a substrate, and a cofactor or inhibitor. The binding of the substrate and cofactor or inhibitor to the enzyme leads to the formation of a stable ternary complex, which can either promote or inhibit enzyme activity depending on the specific reaction and the properties of the molecules involved. For example, in the case of dihydrofolate reductase (DHFR), the enzyme forms a ternary complex with its substrate dihydrofolate (DHF) and the cofactor NADPH during the catalytic cycle. The formation of the DHFR-DHF-NADPH ternary complex is an important step in the reduction of DHF to tetrahydrofolate (THF), and helps to position the reactants in the proper orientation for the transfer of hydride ion from NADPH to DHF.

This is followed by the transfer of a hydride ion from NADPH to the C6 position of DHF, which results in the formation of a highly reactive intermediate. The intermediate then undergoes a series of rearrangements and proton transfers, which ultimately leads to the formation of THF and the release of NADP+. The mechanism of DHFR is highly conserved across species, and is essential for the synthesis of nucleotides and certain amino acids.

Thymidylate kinase (TMPK)

Thymidylate kinase (TMPK) is an enzyme that plays a crucial role in DNA synthesis by catalyzing the transfer of a phosphate group from ATP to thymidine diphosphate (TDP) to produce thymidine triphosphate (TTP), which is an essential building block for DNA. The overall structure of TMPK typically consists of a single polypeptide chain composed of several alpha helices and beta sheets. TMPK enzymes are typically homodimeric, meaning that they are composed of two identical subunits that come together to form an active enzyme. Each subunit contains an ATP-binding domain and a thymidine-binding domain, which are connected by a flexible linker region that allows for conformational changes during catalysis. The active site of TMPK contains a conserved lysine residue that plays a key role in catalysis by interacting with the gamma-phosphate of ATP and facilitating the transfer of the phosphate group to TDP. Other residues, such as arginine and aspartate, also play important roles in substrate binding and catalysis. The structure of TMPK is highly conserved across different organisms, indicating that it has a critical and conserved function in DNA synthesis.

The smallest known version of Thymidylate kinase (TMPK) is the monomeric form found in the bacterium Mycoplasma genitalium, which consists of 146 amino acids. This monomeric form lacks the flexible linker region found in the dimeric form and has a simpler overall structure. However, it still contains the key residues necessary for catalysis and maintains the conserved ATP and thymidine binding domains found in other forms of TMPK. If a cell lacked Thymidylate kinase (TMPK), it would be unable to efficiently synthesize thymidine nucleotides, which are necessary for DNA synthesis and replication. The cell would not be able to maintain proper DNA integrity.  Thymidylate kinase (TMPK) is essential for the viability of most if not all organisms as it plays a critical role in the synthesis of thymidine nucleotides, which are necessary for DNA synthesis and replication. Without TMPK, cells would not be able to efficiently synthesize thymidine nucleotides.

The mechanism of Thymidylate kinase (TMPK) involves the transfer of a phosphate group from ATP to the 5'-hydroxyl group of dTMP, resulting in the formation of dTDP (deoxythymidine diphosphate). This reaction requires the binding of both ATP and dTMP to the enzyme, followed by the transfer of the phosphate group from ATP to dTMP. The reaction is catalyzed by a conserved Asp residue, which serves as a base to deprotonate the 5'-hydroxyl group of dTMP and facilitate the nucleophilic attack of the phosphate group from ATP. The resulting dTDP can then be used as a substrate for DNA synthesis and replication.

Thymidylate kinase (TMPK) depends on the availability of its substrates, ATP and deoxythymidine monophosphate (dTMP), as well as the proper folding of the protein itself. The enzymatic activity of TMPK requires the binding of both substrates to the enzyme active site, which then allows for the transfer of a phosphate group from ATP to dTMP, resulting in the formation of thymidine diphosphate (dTDP) and ADP. Therefore, the proper functioning of TMPK is crucial for the synthesis of DNA and cell growth, as it ensures the availability of thymidine nucleotides, which are essential building blocks for DNA replication and repair.


Nucleoside diphosphate kinase (NDPK)

Nucleoside diphosphate kinase (NDPK) is an enzyme that catalyzes the transfer of a phosphate group from a nucleoside triphosphate (such as ATP) to a nucleoside diphosphate (such as UDP or ADP), producing a nucleoside triphosphate and a nucleoside monophosphate. The reaction is important for maintaining the balance of nucleotide triphosphate and diphosphate pools in the cell, and is also involved in the regulation of various cellular processes, including DNA replication, RNA transcription, and signal transduction. The overall structure of NDPK is characterized by a homohexameric assembly, with each monomer consisting of a central core domain and an N-terminal tail domain. The core domain contains a highly conserved nucleotide-binding site, which is responsible for binding the nucleoside diphosphate substrate and the nucleoside triphosphate donor. The N-terminal tail domain, which varies in length and sequence between different NDPK isoforms, is thought to play a role in regulating enzyme activity and subcellular localization. The hexameric assembly of NDPK is arranged in a head-to-tail fashion, forming a ring-like structure that surrounds a central pore. The nucleoside diphosphate substrate and the nucleoside triphosphate donor enter the pore and bind to the nucleotide-binding site of the core domain, where the transfer of the phosphate group occurs. The transfer reaction is thought to be facilitated by conformational changes in the enzyme that occur upon binding of the nucleotide substrates. Overall, NDPK is a highly conserved enzyme that plays an important role in the maintenance of nucleotide pools and the regulation of cellular processes. Its hexameric structure allows for efficient catalysis and regulation, and its nucleotide-binding site is a common target for drugs and other small molecules that can modulate its activity.

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The pentose phosphate pathway

The pentose phosphate pathway (PPP) is a metabolic pathway that operates parallel to glycolysis and plays a critical role in generating important cellular components and reducing power in the form of NADPH. The pathway starts with glucose-6-phosphate, a six-carbon sugar derived from glucose metabolism. The first two reactions of the pathway involve oxidative steps, which result in the reduction of NADP+ to NADPH and the release of one molecule of CO2. These reactions are catalyzed by the enzymes glucose-6-phosphate dehydrogenase and 6-phosphogluconolactonase. After the oxidative steps, the pathway proceeds through a series of nonoxidative reactions, collectively known as the nonoxidative branch. These reactions involve the rearrangement and interconversion of sugars of various carbon lengths. Through these nonoxidative steps, the pathway generates a range of carbohydrates, including three-, four-, five-, six-, and seven-carbon sugars. Some of these intermediates can enter the glycolytic pathway for further metabolism or serve as precursors for nucleotide synthesis. The enzymes involved in the pentose phosphate pathway are particularly abundant in the cytoplasm of liver and adipose cells. In these tissues, the pathway plays a crucial role in providing NADPH for reductive reactions involved in fatty acid synthesis. NADPH is required as a reducing agent in the synthesis of fatty acids, cholesterol, and other biomolecules. The cytosol, where these enzymes are located, is the site of fatty acid synthesis, and NADPH produced by the pentose phosphate pathway serves as a vital source of reducing power for these biosynthetic reactions.  The pentose phosphate pathway provides an important link between carbohydrate metabolism and the generation of reducing power in the form of NADPH. This pathway is crucial for supporting biosynthetic processes, such as fatty acid synthesis, and plays a role in maintaining redox balance within cells.

The pentose phosphate pathway (PPP), also known as the phosphogluconate pathway, is essential for producing ribose-5-phosphate (R5P), which is an important molecule involved in nucleotide synthesis and various other cellular processes.  R5P is a key precursor for the synthesis of nucleotides, which are the building blocks of DNA and RNA. R5P is converted into 5-phosphoribosyl-1-pyrophosphate (PRPP), which serves as a common precursor for the de novo synthesis of purine and pyrimidine nucleotides.  The PPP generates nicotinamide adenine dinucleotide phosphate (NADPH). NADPH is an essential reducing agent used in numerous cellular processes. It is required for biosynthetic reactions that involve the reduction of various molecules, including fatty acid synthesis, cholesterol synthesis, and nucleotide synthesis. NADPH is also involved in maintaining cellular redox balance and serves as a cofactor for antioxidant defense mechanisms, such as the reduction of reactive oxygen species (ROS).  NADPH produced by the PPP is crucial for the regeneration of glutathione (GSH), a potent antioxidant molecule. GSH helps protect cells from oxidative damage by neutralizing ROS and other harmful reactive species. NADPH is required to maintain an adequate supply of reduced GSH, which is essential for protecting cellular components and maintaining cellular viability under oxidative stress conditions. The PPP is the primary pathway for generating R5P, a five-carbon sugar phosphate. R5P is a critical precursor for nucleotide synthesis, as it contributes to the formation of both purine and pyrimidine nucleotides. Nucleotides are essential for DNA and RNA synthesis, energy metabolism (ATP and NAD+), and various signaling processes. Without the PPP and its ability to produce R5P, the biosynthesis of nucleotides would be severely impaired, hindering cellular growth, replication, and overall viability.  The PPP provides metabolic flexibility to the cell. By branching off from glycolysis, the PPP allows for the production of NADPH and R5P while diverting glucose away from energy production in the form of ATP. This metabolic diversion is particularly important in situations where the cell requires anabolic processes or antioxidant defense more than energy production. The PPP is an indispensable pathway for cellular life. Its functions in NADPH production, antioxidant defense, nucleotide synthesis, and metabolic flexibility are vital for maintaining cellular viability, supporting growth, and adapting to various metabolic demands. The absence or impairment of the PPP would disrupt multiple essential cellular processes, potentially leading to compromised cell function, impaired growth, and increased susceptibility to oxidative damage.

The substrates of the pentose phosphate pathway

The substrates of the pentose phosphate pathway (PPP) are primarily glucose-6-phosphate (G6P) and various sugar phosphates. Following are the key substrates involved in the different reactions of the PPP:

Glucose-6-Phosphate (G6P) is the primary substrate of the pentose phosphate pathway. It is derived from glucose through the action of the enzyme hexokinase or glucokinase, depending on the tissue or cell type. G6P serves as the starting point for both the oxidative and non-oxidative phases of the PPP.

NADP+ (Nicotinamide Adenine Dinucleotide Phosphate) acts as a coenzyme in the oxidative phase of the PPP. It is reduced to NADPH (Nicotinamide Adenine Dinucleotide Phosphate) during the reaction catalyzed by glucose-6-phosphate dehydrogenase (G6PD). NADPH is a crucial product of the PPP and serves as a reducing agent for biosynthetic reactions and antioxidant defense.

Ribulose-5-Phosphate (Ru5P) is an important intermediate formed during the oxidative phase of the PPP. It is generated from the oxidation of G6P by G6PD. Ru5P can be further converted into other sugar phosphates or utilized for nucleotide biosynthesis.

Xylulose-5-Phosphate (Xu5P) is another intermediate produced during the oxidative phase of the PPP. It is derived from Ru5P through a series of enzymatic reactions. Xu5P can be isomerized to ribose-5-phosphate (R5P) or converted into other sugar phosphates.

Ribose-5-Phosphate (R5P) is a key metabolite generated in the non-oxidative phase of the PPP. It is derived from Xu5P or can be directly produced from G6P through a series of enzymatic reactions. R5P is a crucial precursor for nucleotide synthesis, as well as the synthesis of coenzymes and other important biomolecules.

These substrates play essential roles in the PPP, facilitating the generation of NADPH, producing important metabolic intermediates, and providing building blocks for nucleotide and coenzyme biosynthesis.

Synthesis of NADP+ (Nicotinamide Adenine Dinucleotide Phosphate)

NADP+ is synthesized before NADPH is. Such a reaction usually starts with NAD+. The synthesis of NADP+ and NADPH involves several enzymatic reactions and pathways within the cell.  In the de novo pathway, NADP+ is synthesized from simple molecules through a series of enzymatic reactions. The starting point is the amino acid tryptophan, which undergoes a multistep process to form quinolinic acid. Quinolinic acid is then converted to nicotinic acid mononucleotide (NaMN) by the enzyme quinolinate phosphoribosyltransferase. NaMN is subsequently adenylated by NaMN adenylyltransferase to form nicotinic acid adenine dinucleotide (NaAD). Finally, NaAD is phosphorylated by NAD+ kinase to produce NADP+.

The salvage pathway utilizes preformed molecules, such as nicotinamide and nicotinic acid, to synthesize NADP+. Nicotinamide can be converted to nicotinamide mononucleotide (NMN) by the enzyme nicotinamide phosphoribosyltransferase (NAMPT). NMN is then adenylated by NMN adenylyltransferase to form nicotinic acid mononucleotide (NaMN), which is further converted to NADP+ by NAD+ kinase. Alternatively, nicotinic acid can be converted to NaMN directly by nicotinic acid phosphoribosyltransferase (NAPT), followed by the same steps as described above.

NAD+ kinase is an enzyme that plays a crucial role in the synthesis of NADP+. It phosphorylates NAD+ to generate NADP+ by adding an extra phosphate group. This enzyme can accept NAD+ from either the de novo pathway or the salvage pathway and convert it to NADP+. Notably, some forms of NAD+ kinase, particularly the one found in mitochondria, can also accept NADH and directly convert it into NADPH.  ADP-ribosyl cyclase is an enzyme involved in the salvage pathway that allows for the synthesis of NADP+ from nicotinamide. It catalyzes the conversion of nicotinamide to nicotinic acid mononucleotide (NaMN), which serves as an intermediate in the synthesis of NADP+.  NADP+ phosphatase is an enzyme responsible for the conversion of NADPH back to NADH. This enzyme helps maintain a balance between NADP+ and NADPH levels in the cell.

The oxidative phase and the non-oxidative phase.

The oxidative phase primarily generates NADPH, which is important for reductive biosynthesis and oxidative stress defense, while the non-oxidative phase allows for the generation of important metabolic intermediates needed for various cellular processes. The division of the PPP into these two phases enables cells to efficiently meet their requirements for both reducing power (NADPH) and important metabolic building blocks.



In the pentose phosphate pathway (PPP), there are two distinct phases: the oxidative phase and the non-oxidative phase. 
The oxidative phase of the pentose phosphate pathway (PPP), also known as the phosphogluconate pathway or hexose monophosphate pathway, is responsible for the oxidation and decarboxylation of glucose 6-phosphate (G6P) at carbon-1. This phase generates NADPH and pentose phosphates, which have important roles in cellular metabolism. The primary function of the oxidative phase is to produce NADPH, a crucial reducing agent used in various biosynthetic reactions. NADPH provides the necessary reducing power for processes such as fatty acid and cholesterol synthesis, as well as the synthesis of steroid hormones. These reactions occur in rapidly growing tissues or tissues with high demands for biosynthetic processes. By channeling more G6P through the oxidative phase of the PPP, these tissues can generate an ample supply of NADPH to support their biosynthetic activities. Additionally, the oxidative phase of the PPP produces pentose phosphates, most notably ribose 5-phosphate. Ribose 5-phosphate serves as a precursor for nucleotide and nucleic acid synthesis. Rapidly dividing cells, including those found in tissues such as bone marrow, skin, and intestinal mucosa, require ribose 5-phosphate for the production of RNA, DNA, and various coenzymes involved in energy metabolism and cellular signaling. The first phase of the pentose phosphate pathway involves two oxidative reactions. In the initial step, G6P undergoes oxidation and decarboxylation, catalyzed by the enzyme glucose 6-phosphate dehydrogenase (G6PD). This reaction generates NADPH and converts G6P into 6-phosphoglucono-δ-lactone. The lactone is then hydrolyzed by the enzyme lactonase, resulting in the formation of 6-phosphogluconate. In the subsequent step, 6-phosphogluconate is further oxidized by the enzyme 6-phosphogluconate dehydrogenase, generating another molecule of NADPH and converting 6-phosphogluconate into ribulose 5-phosphate. This completes the oxidative phase of the pathway and results in the production of NADPH and pentose phosphates.

The second phase of the pentose phosphate pathway involves nonoxidative steps that interconvert various pentose phosphates and regenerate G6P. These steps allow for the recycling of pentose phosphates and the conversion of multiple molecules of pentose phosphates back into glucose 6-phosphate. This recycling process ensures a continuous supply of NADPH and provides flexibility in utilizing the pentose phosphates for nucleotide synthesis or as a source of energy through the glycolytic pathway.



The role of NADPH in the reductive steps of the PPP refers to its involvement in the oxidative phase of the pathway. During the oxidative phase, a series of enzymatic reactions occur, leading to the production of NADPH. Here's a breakdown of the reductive steps and the role of NADPH: Oxidation of Glucose-6-Phosphate: In the first step of the oxidative phase, glucose-6-phosphate (G6P) is oxidized by the enzyme glucose-6-phosphate dehydrogenase (G6PD). This reaction results in the production of 6-phosphoglucono-delta-lactone and NADPH. The NADPH generated in this step acts as a reducing agent. Conversion of 6-Phosphoglucono-delta-lactone: The 6-phosphoglucono-delta-lactone produced in the previous step is converted into 6-phosphogluconate by the enzyme lactonase. Decarboxylation and Dehydrogenation: 6-Phosphogluconate is then decarboxylated and dehydrogenated by the enzyme 6-phosphogluconate dehydrogenase (6PGD). This reaction generates ribulose-5-phosphate (Ru5P), NADPH, and carbon dioxide. The NADPH produced during the oxidative phase of the PPP is crucial for providing reducing equivalents in various cellular processes. NADPH is utilized in anabolic pathways such as fatty acid synthesis, cholesterol synthesis, and nucleotide synthesis. It acts as a reducing agent, donating electrons to facilitate the reduction of other molecules. The reductive steps of the PPP refer to the generation of NADPH through a series of enzymatic reactions in the oxidative phase of the pathway. NADPH serves as a critical cofactor for reductive biosynthetic reactions and is essential for maintaining cellular redox balance and providing the necessary reducing power for numerous metabolic processes.  The PPP helps maintain redox balance within the cell. It provides an alternative pathway for glucose metabolism, diverting glucose-6-phosphate away from glycolysis. By producing NADPH, the PPP helps replenish the reducing equivalents needed to counteract oxidative stress and maintain cellular redox homeostasis. The oxidative phase of the PPP generates ribulose-5-phosphate (Ru5P), which can be isomerized to produce R5P. R5P is used in the synthesis of nucleotides, coenzymes (such as ATP, NAD+, and FAD), and other molecules involved in cell proliferation and growth.

The pentose phosphate pathway (PPP) involves a series of enzymatic reactions. The number of enzymes involved may vary depending on the specific organism and the variations in the pathway. Here is a list of the enzymes commonly associated with the PPP:

1. Glucose-6-phosphate dehydrogenase (G6PD) catalyzes the conversion of glucose-6-phosphate (G6P) to 6-phosphoglucono-delta-lactone, generating NADPH in the process.
2. Lactonase: Converts 6-phosphoglucono-delta-lactone to 6-phosphogluconate.
3. 6-Phosphogluconate dehydrogenase (6PGD): Catalyzes the decarboxylation and dehydrogenation of 6-phosphogluconate, producing ribulose-5-phosphate (Ru5P), NADPH, and carbon dioxide.
4. Transketolase: Transfers a two-carbon fragment from a ketose sugar to an aldose sugar, facilitating the interconversion between various sugar phosphates in the non-oxidative phase of the PPP.
5. Transaldolase: Catalyzes the transfer of a three-carbon fragment between sugar phosphates in the non-oxidative phase of the PPP.

These five enzymes represent the core reactions of the PPP. However, it's important to note that there may be additional enzymes or variations in different organisms or under specific physiological conditions. Some organisms may possess alternative or bypass pathways that can affect the overall flux through the PPP. Variations in the PPP can occur across different domains of life. While the core reactions and enzymes involved are generally conserved, there can be differences in regulatory mechanisms, isoforms of enzymes, or the presence of additional enzymes that modify the pathway's flux or functions. For example, some microorganisms have unique enzymes or alternative pathways that bypass certain steps of the PPP.



The pentose phosphate pathway is a metabolic pathway that operates parallel to glycolysis and plays a crucial role in the metabolism of glucose. It has two distinct phases: the oxidative phase and the nonoxidative phase.
During the oxidative phase, glucose 6-phosphate (G6P) is oxidized and decarboxylated to produce two important products: NADPH and ribose 5-phosphate (R5P). NADPH is a reducing agent that is involved in numerous cellular processes, including the synthesis of fatty acids, cholesterol, and steroids, as well as the detoxification of reactive oxygen species (ROS) through the reduction of glutathione disulfide (GSSG) to its reduced form (GSH). This process helps protect cells from oxidative damage. R5P, on the other hand, serves as a precursor for the biosynthesis of nucleotides (the building blocks of DNA and RNA), coenzymes (such as NAD+ and FAD), and other important biomolecules involved in cell growth and proliferation. In cells that do not require R5P for biosynthesis, the nonoxidative phase of the pentose phosphate pathway comes into play. This phase involves a series of reversible reactions that convert the intermediates of the pathway back to glycolytic intermediates. Specifically, six molecules of the pentose sugars generated in the oxidative phase are rearranged and converted into five molecules of G6P. This recycling process allows for the continuous production of NADPH, which is important for maintaining the cell's redox balance and supporting various biosynthetic reactions. Additionally, during the nonoxidative phase, the pathway can also generate other sugar phosphates, such as fructose 6-phosphate and glyceraldehyde 3-phosphate, which can be further metabolized through glycolysis or used in other biosynthetic pathways.


The pentose phosphate pathway. The numerals in the blue circles indicate the steps discussed in the text.

In most animal tissues, glucose 6-phosphate (G6P) is primarily catabolized through glycolysis, where it is converted to pyruvate, which is further oxidized via the citric acid cycle to generate ATP. However, G6P can also undergo alternative catabolic pathways, such as the pentose phosphate pathway (PPP) or phosphogluconate pathway, which play specialized roles in certain tissues. The PPP is an oxidative pathway that utilizes NADP+ as the electron acceptor, leading to the production of NADPH. NADPH is a crucial reducing agent required for reductive biosynthetic reactions and for counteracting the damaging effects of reactive oxygen species (ROS). Certain tissues with high proliferative rates, including bone marrow, skin, intestinal mucosa, and tumors, rely on the PPP to generate ribose 5-phosphate, a pentose sugar used in the synthesis of RNA, DNA, and coenzymes like ATP, NADH, FADH2, and coenzyme A. The PPP supplies these rapidly dividing cells with the necessary building blocks for nucleic acid synthesis and energy production. In other tissues, the primary importance of the PPP lies in the generation of NADPH rather than pentose sugars. Tissues involved in extensive fatty acid synthesis (such as the liver, adipose tissue, and lactating mammary gland) or active synthesis of cholesterol and steroid hormones (such as the liver, adrenal glands, and gonads) require a significant amount of NADPH. NADPH serves as a reducing equivalent in the biosynthesis of fatty acids and cholesterol and plays a crucial role in maintaining redox balance within these tissues. Additionally, cells that are directly exposed to oxygen, such as erythrocytes (red blood cells) and the cells of the lens and cornea, rely on NADPH to prevent oxidative damage caused by reactive oxygen species. By maintaining a high ratio of NADPH to NADP+ and a high ratio of reduced to oxidized glutathione, these cells create a reducing environment that protects proteins, lipids, and other sensitive molecules from oxidative damage. In the case of red blood cells,  erythrocytes, the importance of NADPH production through the PPP is underscored by the fact that a genetic defect in the first enzyme of the pathway, glucose 6-phosphate dehydrogenase (G6PD), can have severe medical consequences. G6PD deficiency can lead to hemolytic anemia, a condition characterized by the destruction of red blood cells due to their increased susceptibility to oxidative stress. The PPP's role in generating NADPH becomes crucial in preventing oxidative damage and maintaining the integrity and function of erythrocytes.

1. Oxidation of glucose-6-phosphate

The oxidation of glucose-6-phosphate in the pentose phosphate pathway is catalyzed by the enzyme glucose-6-phosphate dehydrogenase (G6PDH). This reaction leads to the formation of a cyclic ester called the lactone of phosphogluconic acid and the production of NADPH. G6PDH is highly specific for NADP+ as its cofactor, and it plays a crucial role as the first step of the pentose phosphate pathway. This reaction is considered irreversible and is tightly regulated to maintain metabolic balance. The specificity for NADP+ ensures that NADPH, the reduced form of NADP+, is generated, which is essential for various reductive reactions within the cell. The regulation of G6PDH is complex and involves multiple factors. One important regulatory mechanism is feedback inhibition. The enzyme is strongly inhibited by its product, NADPH. When NADPH levels are high, it acts as a negative feedback signal, effectively slowing down the activity of G6PDH to prevent excessive production of NADPH. Additionally, G6PDH is also inhibited by fatty acid esters of coenzyme A, which are intermediates of fatty acid biosynthesis. This inhibition helps to coordinate the metabolic flux between the pentose phosphate pathway and fatty acid synthesis. The inhibition of G6PDH by NADPH is dependent on the cytosolic NADP+/NADPH ratio. The regulation of G6PDH activity is essential for maintaining the balance between NADP+ and NADPH levels in the cell. It ensures that NADPH is produced when needed for reductive reactions, while avoiding excessive accumulation that could disrupt cellular redox homeostasis.

Glucose-6-phosphate dehydrogenase (G6PDH)

G6P is a six-carbon sugar in the form of a cyclic hemiacetal, with the carbon at position 1 (C1) in the aldehyde oxidation state. The G6PD enzyme facilitates the oxidation of G6P, resulting in the formation of a cyclic ester known as 6-phosphoglucono-lactone. This reaction involves the transfer of a hydride ion (H-) from C1 of G6P to the NADP+ cofactor, leading to the reduction of NADP+ to NADPH. The enzyme G6PD is highly specific for NADP+ as its cofactor, meaning it selectively uses NADP+ in the oxidation-reduction reaction. This specificity ensures the generation of NADPH, which serves as a crucial reducing agent in various metabolic pathways. One interesting aspect of G6PD regulation is the strong inhibition by NADPH, the reduced form of NADP+. When NADPH levels are elevated, it acts as a feedback inhibitor of G6PD. This inhibition helps maintain the balance of NADP+/NADPH ratios in the cell and prevents an excessive production of NADPH when it is not required.

Glucose-6-phosphate dehydrogenase (G6PDH) is a monomeric enzyme.  G6PDH catalyzes the oxidation of G6P, converting it into 6-phosphoglucono-lactone and producing NADPH as a reduced cofactor. The average size of G6PDH in terms of amino acids varies among different species. In humans, it consists of approximately 515 amino acids. G6PDH contains a metal co-factor in its reaction pocket, specifically a zinc ion (Zn2+). The metal co-factor is essential for the proper folding and stability of the enzyme and for facilitating its catalytic activity.  Apart from the metal co-factor, G6PDH requires NADP+ as a cofactor for the oxidation-reduction reaction. The mechanism of G6PDH involves the binding of G6P and NADP+ to the enzyme's active site. G6P undergoes oxidation, resulting in the transfer of a hydride ion to NADP+ and the formation of NADPH. This process involves a series of chemical reactions and enzyme-substrate interactions. Here is an elucidation of the process:

Enzyme structure

The structure and function of glucose-6-phosphate dehydrogenase (G6PD) exhibit remarkable precision and fine-tuning to enable the enzyme to catalyze its specific reaction with specificity and error proneness. G6PD is typically found as a dimer composed of two identical monomers, and under certain conditions, these dimers can further dimerize to form tetramers. Each monomer within the complex possesses a substrate binding site that interacts with glucose-6-phosphate (G6P) and a catalytic coenzyme binding site that accommodates NADP+/NADPH using the Rossman fold. Additionally, some organisms, including humans, have an extra NADP+ binding site known as the NADP+ structural site, the exact purpose remains unknown. Several regions in G6PD exhibit functional and structural conservation across different organisms. These include a 9-residue peptide sequence (RIDHYLGKE), located in the substrate binding site, a nucleotide-binding fingerprint (GxxGDLA), and a partially conserved sequence (EKPxG) near the substrate binding site. These conserved regions are crucial for the proper functioning of the enzyme and play a role in substrate recognition and binding.

The presence of functional and structurally conserved regions in G6PD across different organisms strongly suggests that the enzyme was created fully functional from its inception. These conserved regions, such as the 9-residue peptide sequence (RIDHYLGKE), the nucleotide-binding fingerprint (GxxGDLA), and the partially conserved sequence (EKPxG), are crucial for the enzyme's proper functioning and play specific roles in substrate recognition and binding. If G6PD were to emerge gradually through a step-by-step process, it would involve the accumulation of multiple genetic changes over an extended period. However, the development of an enzyme with functional activity through such incremental changes is highly unlikely. In a stepwise process, intermediates would be formed during the gradual accumulation of genetic changes. However, these intermediates would not possess the precise arrangement and coordination of functional regions required for the proper functioning of G6PD. Without the conserved regions that are crucial for substrate recognition and binding, the intermediates would lack the ability to catalyze the specific reaction efficiently. Moreover, the conserved regions in G6PD exhibit intricate molecular interactions and precise positioning of residues. These features are not easily attainable through a step-by-step process, as each incremental change would likely disrupt the existing functionality or render the intermediate non-functional. The simultaneous occurrence of multiple genetic changes to generate the conserved regions and their specific interactions would require a coordinated and purposeful design rather than a gradual, random process. This strongly suggests that the enzyme's fully functional state was present from its origin. The intricate and specific arrangement of these conserved regions points towards intelligent design rather than a series of random, non-functional intermediates that would be expected from a stepwise evolutionary process.

The crystal structure of G6PD reveals an extensive network of electrostatic interactions and hydrogen bonding involving G6P, water molecules, lysines, arginine, histidines, glutamic acids, and other polar amino acids. These interactions contribute to the precise positioning of the substrate and facilitate the catalytic activity of the enzyme. The proline residue at position 172 is particularly noteworthy as it plays a crucial role in positioning the lysine residue at position 171 correctly with respect to the substrate, G6P. The presence of proline at position 172 influences the conformation of the enzyme and ensures the proper alignment of key functional residues. Mutations in G6PD can lead to enzymopathy and deficient G6PD activity. Interestingly, mutations causing disease are frequently found near the NADP+ structural site. Access to crystal structures has allowed scientists to model the structures of various mutants, providing insights into the impact of specific mutations on the enzyme's function. Mutations in critical regions near the NADP+ binding site, G6P binding site, or the interface between monomers can occur without completely disrupting the enzyme's function. This suggests that the overall structure and function of G6PD are finely tuned and can tolerate specific mutations while still maintaining enzymatic activity. The precise adjustments and tuning observed in G6PD are essential for the enzyme's specific reaction and error proneness. The conserved regions, an extensive network of interactions, and proper positioning of critical residues ensure the specific recognition and binding of the substrate, G6P. These structural features, along with the flexibility to accommodate certain mutations, contribute to the enzyme's ability to carry out its function accurately.

If the residues at positions 172 and 171 in G6PD were not the correct ones or if they were altered due to mutations, it would likely have a significant impact on the enzyme's structure and function. The proline residue at position 172 plays a crucial role in positioning the lysine residue at position 171 correctly with respect to the substrate, G6P. This proper alignment is essential for the enzymatic reaction to occur accurately. If the proline residue is absent or replaced with a different amino acid, it could disrupt the conformation of the enzyme and result in the misalignment of key functional residues. The misalignment of residues at positions 171 and 172 would likely lead to a loss of the enzyme's ability to properly interact with the substrate, G6P. This could affect the binding affinity and specificity of G6PD for G6P, impairing the catalytic activity of the enzyme. As a result, the enzyme may exhibit reduced or even complete loss of enzymatic function. Mutations in G6PD, particularly those near the NADP+ structural site, can lead to enzymopathy and deficient G6PD activity. This indicates the critical role of these regions in maintaining the enzyme's stability and function. Mutations that affect the proper alignment of residues, including those at positions 171 and 172, can disrupt the overall structure and compromise the enzyme's ability to catalyze the reaction effectively. It is highly unlikely that a random process would generate the correct arrangement of these residues to achieve the desired enzymatic activity. This is because the correct positioning of residues requires specific interactions, such as electrostatic interactions and hydrogen bonding, as well as precise conformational changes to accommodate the substrate. The extensive network of interactions observed in the crystal structure of G6PD, involving G6P, water molecules, lysines, arginine, histidines, glutamic acids, and other polar amino acids, indicates a highly specialized and finely tuned system. These interactions contribute to the precise positioning of the substrate and facilitate the catalytic activity of the enzyme. The presence of conserved regions and the specific alignment of residues further highlight the design-like nature of G6PD. If the residues at positions 172 and 171 were randomly positioned or altered through a random process, it would likely result in a loss of proper alignment and impaired enzymatic function. Achieving the correct alignment of functional residues requires a specific sequence of amino acids and their precise three-dimensional arrangement, which is highly unlikely to occur by chance alone. Furthermore, mutations near the NADP+ structural site, which frequently lead to disease-causing effects, highlight the sensitivity of the enzyme's structure and function to changes in critical regions. These observations suggest that G6PD is finely tuned and optimized for its specific function, further supporting the idea of a designed set-up.

The NADP+ structural site

The NADP+ structural site refers to a specific region within the enzyme where the molecule NADP+ binds. This site is distinct from the catalytic coenzyme NADP+ binding site and serves a different purpose in the enzyme's function.
The structural site is located at a considerable distance, greater than 20Å, away from both the substrate binding site and the catalytic coenzyme NADP+ binding site.  It has been observed that the presence of NADP+ at the structural site promotes the dimerization of dimers, leading to the formation of enzyme tetramers.  The NADP+ structural site exhibits specific characteristics and interactions that contribute to its stability and long-term functionality. It contains the nucleotide-binding fingerprint, a distinct sequence motif involved in binding nucleotide molecules. The structural site is characterized by a strong network of hydrogen bonding, involving electrostatic charges that are distributed across multiple atoms through hydrogen bonds with four water molecules. Additionally, there are robust hydrophobic stacking interactions that result in overlapping π systems, further enhancing the stability of NADP+ binding to the structural site. Mutations near the NADP+ structural site have been found to have significant implications for the stability of the enzyme. Over 40 severe class I mutations have been identified in this region, affecting the long-term stability of G6PD and leading to G6PD deficiency.  

High specificity

6-phosphate dehydrogenase is highly specific for NADP 1; the K M for NAD 1 is about a thousand times as great as that for NADP 1.  In enzymology, K M (Michaelis constant) is a measure of the substrate concentration at which an enzyme reaches half of its maximum reaction rate (Vmax). It is a fundamental parameter used to characterize the affinity of an enzyme for its substrate. Specifically, K M represents the substrate concentration at which the rate of the enzymatic reaction is equal to half of Vmax. A lower K M value indicates a higher affinity of the enzyme for its substrate, meaning that the enzyme can effectively bind to and catalyze the reaction even at lower substrate concentrations. Conversely, a higher K M value indicates a lower affinity of the enzyme for its substrate, requiring higher substrate concentrations to achieve half of the maximum reaction rate. In the case of glucose-6-phosphate dehydrogenase (G6PD), the K M for NAD+ (oxidized form) is about a thousand times greater than the K M for NADP+ (phosphorylated form). This implies that G6PD has a significantly higher affinity for NADP+ compared to NAD+. It means that G6PD can efficiently utilize NADP+ as a coenzyme in its catalytic reaction, while NAD+ would need to be present at much higher concentrations to achieve the same level of enzymatic activity. If the G6PD did not have a high affinity for NADP+ and instead had a lower affinity or no affinity for it, several consequences could arise: The catalytic efficiency of G6PD would be significantly reduced. The enzyme would struggle to effectively bind and utilize NADP+ as a coenzyme, resulting in a slower rate of enzymatic reaction. This could lead to a decreased production of NADPH, which is crucial for various cellular processes.  G6PD plays a key role in the oxidative phase of the pentose phosphate pathway, where it generates NADPH. A lower affinity for NADP+ would hinder the enzyme's ability to generate NADPH efficiently. This could disrupt the balance of cellular redox reactions and impair the proper functioning of the pentose phosphate pathway. NADPH produced by G6PD is a vital reducing agent for antioxidant defense systems, such as the maintenance of the reduced glutathione pool. With a reduced ability to generate NADPH, cells would be less capable of combating oxidative stress and protecting against damage caused by reactive oxygen species. The reduced production of NADPH could have broader implications for cellular metabolism and homeostasis. NADPH is involved in various biosynthetic processes, including fatty acid and cholesterol synthesis. Insufficient NADPH levels may compromise these pathways and result in metabolic imbalances and cellular dysfunction.

The high specificity of G6PD for NADP+ is evidence of an optimized and finely tuned system. The precise fit between the enzyme and its preferred coenzyme suggests that the system has been designed to efficiently carry out specific functions within the cell. This level of specificity is required to meet the specific functional needs of the cell.  The high affinity of G6PD for NADP+ allows for efficient catalysis of the oxidative reactions in the pathway. A tightly bound coenzyme ensures that the reaction proceeds at a sufficient rate even at lower substrate concentrations. This optimized efficiency suggests that the enzyme has been precisely tuned to maximize its catalytic potential, indicating the presence of design principles. The specific interactions between G6PD and NADP+ involve precise molecular recognition and complementarity. The active site of G6PD is structured to accommodate the unique chemical properties and structure of NADP+. This level of precision in molecular recognition and binding suggests a purposeful arrangement, consistent with a design perspective. The high specificity of G6PD for NADP+ requires specific amino acid residues and structural features within the enzyme. Achieving such specificity would necessitate complex and precise arrangements of these elements. The level of information content required to encode this specificity is indicative of a deliberate design rather than a random, unguided process. The specific fit between G6PD and NADP+ is an example of irreducible complexity. If any key elements or interactions were altered, the enzyme's efficiency and specificity would be compromised, leading to dysfunctional or less optimal metabolic pathways. This implies that the system requires all its components in their precise arrangement from the outset, supporting the notion of intentional design.

Catalytic reaction

G6P initially binds to the active site of G6PDH through specific interactions between the enzyme and substrate molecules. Here are some of the key interactions: G6P contains several hydroxyl groups (-OH), which can form hydrogen bonds with specific amino acid residues present in the active site of G6PDH. These hydrogen bonds help stabilize the binding of G6P to the enzyme. For example, the hydroxyl groups of G6P can interact with amino acid residues such as serine, threonine, or asparagine through hydrogen bonding. The active site of G6PDH may contain charged amino acid residues such as arginine or glutamate. These charged residues can interact with the charged functional groups present in G6P, such as phosphate groups or the aldehyde group. Electrostatic interactions play a role in the specific binding of G6P to the active site of G6PDH.  G6P and the active site of G6PDH also interact through van der Waals forces, which are weak attractive forces between atoms or molecules. Van der Waals interactions occur due to the fluctuating electron distributions within molecules, creating temporary positive and negative charges. These interactions contribute to the overall stability of the G6P-G6PDH complex. The active site of G6PDH has a specific shape that complements the shape of G6P. This shape complementarity allows for optimal binding and interaction between the enzyme and substrate. The active site may have pockets or grooves that accommodate the specific structural features of G6P, ensuring a tight and specific binding. These specific interactions between G6P and G6PDH are crucial for the recognition and binding of the substrate to the enzyme's active site. They provide a specific and precise binding mechanism, ensuring that G6P is properly positioned for the subsequent catalytic reactions to occur.



The active site of G6PDH exhibits remarkable specificity and precision in its interaction with G6P. The shape complementarity, hydrogen bonding, electrostatic interactions, and van der Waals forces are all finely tuned to ensure a tight and specific binding between the enzyme and substrate. Achieving such high specificity and precision requires intricate molecular recognition and precise coordination. The active site of G6PDH is designed to accommodate G6P and facilitate its oxidation and subsequent conversion to 6-phosphogluconolactone. The specific arrangement of amino acid residues in the active site, along with the presence of metal cofactors if applicable, is crucial for the catalytic activity of the enzyme. This is a high level of functional optimization to ensure efficient and accurate enzymatic activity.  The overall structure and function of G6PDH involve a complex interplay of various components, including the active site, cofactors, and other structural elements. Naturalistic unguided processes are highly unlikely to produce such intricate and integrated molecular machinery.  G6PDH is found across different organisms and exhibits a high degree of structural and functional conservation. This conservation suggests that the enzyme's design is essential for its biological function. Considering these factors, it becomes increasingly implausible to attribute the precise and functional design of G6PDH solely to naturalistic unguided processes. The complex and purposeful arrangement of the enzyme's active site and its specific interactions with G6P strongly suggest the involvement of intelligent design in its origin.

G6PDH facilitates the removal of two hydrogen atoms from G6P. This removal is achieved through a series of chemical reactions that involve the transfer of hydride ions (H-) from the C1 carbon of G6P to the enzyme and subsequent transfer to the coenzyme NADP+. Here is an elucidation of the steps involved:  G6PDH catalyzes the initial step by oxidizing G6P. In this process, two hydrogen atoms (H) are removed from the C1 carbon of G6P. The removal of these hydrogen atoms results in the formation of a double bond between C1 and C2, leading to the formation of 6-phosphoglucono-delta-lactone.  The hydrogen atoms removed from G6P in the previous step are transferred to the enzyme G6PDH. Specifically, a hydride ion (H-) is transferred from the C1 carbon of G6P to the enzyme, where it interacts with specific active site residues.  Following the transfer of the hydride ion to the enzyme, the next step involves the transfer of the hydride ion from the enzyme to the coenzyme NADP+. The NADP+ molecule accepts the hydride ion, resulting in the reduction of NADP+ to NADPH. Overall, the series of chemical reactions involve the oxidation of G6P, the transfer of hydride ions from G6P to the enzyme, and the subsequent transfer of the hydride ions from the enzyme to NADP+. The precise details of the chemical reactions and the specific residues involved in the hydride ion transfer may vary depending on the specific isoform of G6PDH and the organism in question. However, the general concept of oxidation, hydride ion transfer, and reduction of NADP+ to NADPH remains consistent.
The efficient and specific catalysis of these reactions by G6PDH ensures the production of NADPH, which is essential for various cellular functions. 

The transferred hydride ions from G6P are temporarily held within the active site of G6PDH. During this step, an intermediate compound is formed, which is an unstable phosphorylated compound.  In the next step, the intermediate compound undergoes spontaneous intramolecular rearrangement, leading to the release of carbon dioxide (CO2). This step is crucial for the conversion of G6P into 6-phosphoglucono-lactone.  After the release of CO2, the hydride ion is transferred from the enzyme's active site to NADP+. This transfer results in the reduction of NADP+ to NADPH.  The final step involves the release of the cyclic ester, 6-phosphoglucono-lactone, from the active site of G6PDH. Simultaneously, NADPH is released as a product of the reaction. This series of reactions catalyzed by G6PDH leads to the oxidation of G6P, the transfer of a hydride ion to NADP+, and the formation of NADPH. The role of G6PDH is to facilitate the efficient transfer of electrons and hydride ions, enabling the generation of NADPH, which is essential for various cellular processes, including reductive reactions and antioxidant defense.

The intricate series of chemical reactions facilitated by glucose-6-phosphate dehydrogenase (G6PDH) are a goal-oriented process. The precise coordination and interplay of multiple steps, each with specific functions, are evidence of a purposefully designed implementation to achieve a specific goal efficiently.  The overall process of G6P oxidation and NADPH production requires a conceptualization of the desired outcome. It involves the recognition that G6P can serve as a potential energy source and the understanding that its oxidation and NADPH production would be beneficial for cellular functions such as reductive reactions and antioxidant defense.  The design of G6PDH involves the precise arrangement of its active site residues, metal cofactors, and structural elements to facilitate the necessary chemical reactions. The active site is meticulously shaped and positioned to accommodate the specific binding of G6P, ensuring proper orientation and interaction for subsequent steps.  The series of chemical reactions involving G6P oxidation and the transfer of hydride ions to NADP+ occurs in multiple steps. This multi-step process allows for fine control and accuracy, minimizing the potential for errors or side reactions. Each step serves a specific purpose, ensuring the orderly progression of the reactions and the generation of NADPH. The enzymatic process carried out by G6PDH is aimed at achieving the specific goal of NADPH production. The precise interactions, shape complementarity, and molecular recognition between the enzyme and its substrate are crucial for the efficient catalysis of the reactions. This precision ensures that G6P is properly positioned and processed, leading to the desired outcome of NADPH generation.  The involvement of multiple steps also introduces the potential for errors or deviations from the desired outcome. However, the remarkable specificity and efficiency of G6PDH in catalyzing these reactions minimize the occurrence of errors and maximize the production of NADPH, which is essential for various cellular processes. The intricate and purposeful nature of the enzyme's design, the multiple steps involved, and the precision required to achieve the specific goal of NADPH production strongly suggest the involvement of intelligent design. The complexity and functional optimization observed in G6PDH's structure and catalytic mechanism is indicative of an intentional and goal-oriented process, surpassing the likelihood of naturalistic unguided mechanisms to produce such a sophisticated system. G6PDH is regulated by various factors, including feedback inhibition by NADPH. Elevated levels of NADPH act as a negative feedback signal, inhibiting the activity of G6PDH to prevent excessive NADPH production. The regulation of G6PDH activity is also influenced by the NADP+/NADPH ratio in the cytosol.

Regulation, essential for the cell's survival

The regulation of G6PD activity is essential to maintain the appropriate balance of NADPH/NADP+ in the cell.  Maintaining the appropriate balance of NADPH/NADP+ is essential for the cell's proper functioning and survival. NADPH serves as a critical cofactor in numerous essential cellular processes, particularly those involved in combating oxidative stress and supporting biosynthesis. If the balance of NADPH/NADP+ is disrupted and the levels of NADPH become insufficient relative to NADP+, several detrimental consequences can occur:  NADPH is a key component in the cellular antioxidant defense system. It is required to regenerate reduced glutathione (GSH), an important antioxidant molecule that protects cells from oxidative damage. Without sufficient NADPH, the capacity to maintain an optimal antioxidant defense becomes compromised, leading to increased oxidative stress and potential damage to cellular components, including proteins, lipids, and DNA.  NADPH plays a vital role in biosynthetic pathways, such as fatty acid and cholesterol synthesis, nucleotide synthesis, and the production of various macromolecules. These processes require NADPH as a reducing agent for the synthesis of building blocks and energy storage. Insufficient NADPH levels can hamper these biosynthetic pathways, leading to impaired cell growth, energy production, and the synthesis of essential molecules. Without adequate levels of NADPH, cells become more vulnerable to oxidative damage. Reactive oxygen species (ROS), generated during normal cellular metabolism or under stressful conditions, can accumulate and overwhelm the cell's antioxidant defense system. Insufficient NADPH limits the cell's ability to neutralize ROS and counteract oxidative stress, which can result in cellular dysfunction, DNA damage, and even cell death. NADPH and NADP+ are key components of the cellular redox system, maintaining the balance between oxidized and reduced forms of molecules. Disruptions in the NADPH/NADP+ balance can disturb redox homeostasis, affecting numerous redox-sensitive processes and signaling pathways within the cell. This can lead to dysregulation of cellular functions and contribute to various pathological conditions.

G6PD is stimulated by its substrate, G6P, which promotes the enzymatic conversion of G6P to 6-phosphoglucono-δ-lactone. A high ratio of NADPH/NADP+ is required for biosynthetic processes, such as fatty acid synthesis. Increased utilization of NADPH for fatty acid biosynthesis leads to an elevated level of NADP+, which in turn stimulates G6PD to produce more NADPH. This feedback mechanism ensures an adequate supply of NADPH to meet the demands of fatty acid synthesis. In yeast, G6PD is inhibited by long-chain fatty acids, potentially acting as a product inhibition mechanism to regulate fatty acid synthesis that relies on NADPH. Another important regulatory mechanism of G6PD involves acetylation on lysine 403 (Lys403), a conserved residue. Acetylation of Lys403 negatively regulates G6PD by inhibiting the formation of active enzyme dimers and resulting in a complete loss of enzymatic activity. The acetylation of Lys403 sterically hinders the entry of NADP+ into the NADP+ structural site, compromising the stability of the enzyme. Cells have mechanisms to sense extracellular oxidative stimuli and decrease G6PD acetylation in a SIRT2-dependent manner. The deacetylation and activation of G6PD by SIRT2 promote the pentose phosphate pathway, increasing the production of cytosolic NADPH to counteract oxidative damage and protect cells. In addition to posttranslational modifications, regulation of G6PD can also occur through genetic pathways. The isoform G6PDH is regulated at the transcriptional level by transcription factors and posttranscriptional factors. This allows for fine-tuning of G6PD expression and activity in response to specific cellular needs. The intricate regulation of G6PD serves as evidence of a purposeful and sophisticated design. The enzyme is precisely controlled at multiple levels to ensure the proper balance of NADPH/NADP+ and to meet the cellular demands for various processes, including antioxidant defense and biosynthetic pathways. The ability of cells to sense and respond to extracellular oxidative stimuli, such as through SIRT2-mediated deacetylation of G6PD, demonstrates a designed mechanism for adapting to changing environmental conditions. The coordinated regulation of G6PD activity through substrate stimulation, product inhibition, and posttranslational modifications highlights the precision and optimization in the design of the enzyme and its role in cellular metabolism.

2. Lactonase converts 6-phosphoglucono-delta-lactone to 6-phosphogluconate

After the formation of 6-phosphoglucono-γ-lactone in the pentose phosphate pathway, the next step involves the hydrolysis of 6-phosphoglucono-δ-lactone by a specific enzyme called lactonase. This hydrolysis reaction breaks the intramolecular ester bond between the C-1 carboxyl group and the C-5 hydroxyl group, resulting in the formation of 6-phosphogluconate.


Glucose 6-phosphate is oxidized to 6-phosphoglucono-d-lactone to generate one molecule of NADPH. The lactone product is hydrolyzed to 6-phosphogluconate, which is oxidatively decarboxylated to ribulose 5-phosphate with the generation of a second molecule of NADPH.

6-phosphogluconate is a six-carbon sugar acid that serves as an intermediate in the pentose phosphate pathway. It undergoes further enzymatic reactions to produce important metabolic products. One of the key reactions is the oxidative decarboxylation of 6-phosphogluconate catalyzed by the enzyme 6-phosphogluconate dehydrogenase. During oxidative decarboxylation, 6-phosphogluconate dehydrogenase removes a carboxyl group from 6-phosphogluconate, resulting in the formation of ribulose 5-phosphate. This reaction involves the transfer of electrons to an electron acceptor, which in the pentose phosphate pathway is NADP+ (nicotinamide adenine dinucleotide phosphate), yielding NADPH in the process. Ribulose 5-phosphate is an important intermediate in several metabolic pathways. It can be converted into other sugars, such as ribose 5-phosphate, which is a crucial component for nucleotide synthesis and the production of coenzymes like ATP (adenosine triphosphate) and NADH (nicotinamide adenine dinucleotide). Ribulose 5-phosphate is also involved in the synthesis of important molecules like amino acids and certain coenzymes.

Lactonase

The lactonase enzyme is also known as lactonase/acyltransferase and belongs to the family of hydrolases. The primary function of lactonase is to catalyze the hydrolysis of lactones. Lactones are cyclic esters that can be found in various biological systems. The enzyme breaks the ester bond in lactones, converting them into their respective carboxylic acids. Lactonase can exist in both monomeric and multimeric forms. The overall structure of the enzyme is highly variable and can differ depending on the specific lactonase variant. Generally, lactonases are composed of amino acids arranged in a three-dimensional structure. They often possess active sites or binding regions that facilitate their interaction with lactone substrates.  Lactonase exhibits specificity for lactone substrates. The enzyme recognizes and binds to the lactone molecule through specific interactions between amino acid residues in the active site and functional groups present in the lactone structure. Lactonase typically contains one binding site for lactone molecules.  The regulation of lactonase activity can vary depending on the specific variant and the organism it is derived from. Some lactonases can be regulated by allosteric effectors or post-translational modifications, which can modulate their enzymatic activity or expression levels. Lactonases are found in various organisms, including bacteria, plants, and animals. They can exhibit different substrate specificities and can play roles in diverse biological processes, such as quorum sensing regulation in bacteria.

Quorum sensing

Quorum sensing is a mechanism by which bacteria communicate with each other and coordinate their behavior based on changes in population density. It allows bacterial communities to synchronize their activities and act as a collective entity. Quorum sensing is particularly important for bacteria that form biofilms, which are complex, organized communities of bacteria attached to surfaces. In many quorum sensing systems, the signaling molecules involved are AHLs (N-acyl homoserine lactones). AHLs are small molecules that are produced and released by bacteria into their environment. As the bacterial population grows, the concentration of AHLs increases. Once a certain threshold concentration is reached, the bacteria can sense this through AHL receptors and initiate specific cellular responses. Lactonases, specifically AHL lactonases, are enzymes that can degrade and inactivate AHL signaling molecules. They catalyze the hydrolysis of the lactone ring present in AHLs, resulting in the breakdown of AHLs into their corresponding carboxylic acids. By cleaving the lactone ring, lactonases prevent AHLs from binding to their receptors, thus disrupting the quorum-sensing signaling pathway. By modulating the levels of AHLs, lactonases have a direct impact on quorum sensing regulation. They can influence the timing, duration, and intensity of quorum-sensing responses in bacterial populations. This ability to control quorum sensing allows bacteria to coordinate behaviors such as biofilm formation, virulence factor production, antibiotic resistance, and other processes that are advantageous in a community setting. In addition to their role in quorum sensing, lactonases may have other functions in different organisms. For example, in plants, lactonases have been implicated in defense mechanisms against pathogens. They can hydrolyze lactones produced by pathogens and disrupt their signaling pathways, thus interfering with the pathogen's ability to establish an infection.

If lactonase is a multimeric enzyme, the subunits are essential for the proper function of the enzyme. Each subunit contributes to the overall structure and activity of the enzyme, and the absence of any subunit may result in the loss of function.

3. 6-phosphogluconate undergoes oxidation and decarboxylation by 6-phosphogluconate dehydrogenase to form the ketopentose ribulose 5-phosphate

The oxidative decarboxylation of 6-phosphogluconate by 6-phosphogluconate dehydrogenase plays a crucial role in the pentose phosphate pathway. This pathway is involved in the production of NADPH, which is essential for various cellular processes, including biosynthesis of fatty acids and steroids, detoxification of reactive oxygen species, and maintenance of cellular redox balance.

The reaction catalyzed by 6-phosphogluconate dehydrogenase occurs in two distinct steps.

In the first step, NADP1-dependent dehydrogenation takes place. The enzyme 6-phosphogluconate dehydrogenase oxidizes 6-phosphogluconate, utilizing NADP+ as a cofactor. This oxidative reaction results in the removal of two hydrogen atoms from 6-phosphogluconate and their transfer to NADP+, leading to the formation of NADPH. Simultaneously, the conversion of 6-phosphogluconate into a β-keto acid, 3-keto-6-phosphogluconate, occurs. This conversion is crucial as it prepares the substrate for the subsequent decarboxylation step.

In the second step, 3-keto-6-phosphogluconate undergoes decarboxylation. The β-keto acid intermediate is highly susceptible to decarboxylation, meaning that it readily loses a carboxyl group (CO2) from its structure. This decarboxylation reaction is thermodynamically favorable and leads to the formation of d-ribulose-5-phosphate. The release of CO2 allows the molecule to undergo structural rearrangements, resulting in the formation of d-ribulose-5-phosphate.


The 6-phosphogluconate dehydrogenase reaction. Oxidation of the OH group forms an easily decarboxylated beta-keto acid

The resulting d-ribulose-5-phosphate serves as a key substrate for the nonoxidative reactions that occur in the rest of the pentose phosphate pathway. These nonoxidative reactions involve interconversions of various sugar phosphates and play a crucial role in generating important intermediates for nucleotide synthesis, as well as providing precursors for glycolysis and gluconeogenesis.

 6-phosphogluconate dehydrogenase

The structure of 6-phosphogluconate dehydrogenase can vary depending on the organism. It can be either monomeric or multimeric, with each subunit containing a catalytic domain responsible for the enzymatic activity. Notable structural features may include active sites and binding regions involved in substrate recognition and catalysis.  The enzyme specifically acts on 6-phosphogluconate as its substrate. The active site of 6-phosphogluconate dehydrogenase recognizes and binds to 6-phosphogluconate, allowing for the catalytic transformation into d-ribulose-5-phosphate and NADPH. The active site typically consists of amino acid residues that interact with the substrate, facilitating the chemical reaction.  The catalytic activity of 6-phosphogluconate dehydrogenase relies on specific amino acids and groups of atoms within its active site. These include residues that interact with the substrate and facilitate the chemical transformation. The precise arrangement of amino acids, along with their charges, shape, and other molecular features within the active site, is crucial for substrate recognition and binding. The fine-tuning of rotation angles in enzymes can be attributed to intelligent design in the sense that the precise arrangement of amino acids and atoms within the active site allows for optimal substrate recognition and catalysis. It suggests that there is an intentional design to achieve specific functions and enhance the efficiency of enzymatic reactions. This perspective emphasizes the complexity and precision of enzyme structure

Catalytic site

6-Phosphogluconate dehydrogenase (6PGD) is a multimeric enzyme, meaning it consists of multiple subunits that come together to form the active enzyme. Each subunit contains a catalytic site where the enzymatic reactions take place.  The active site of 6PGD is typically located in a cleft or pocket within the enzyme structure, where the catalytic reactions occur. The active site comprises specific amino acid residues that are critical for catalysis. The catalytic mechanism of 6PGD involves the binding of the substrate, 6-phosphogluconate, and the cofactor, NADP+. The active site residues interact with these molecules, facilitating the transfer of hydride ions from 6-phosphogluconate to NADP+, resulting in the formation of NADPH. One important residue in the active site is a histidine residue that acts as a catalytic base, accepting a proton from the substrate during the dehydrogenation reaction. Other residues, such as serine, threonine, and lysine, are involved in stabilizing the substrate and cofactor binding and orienting them for the catalytic reaction. The precise arrangement and interactions of these residues within the active site are crucial for the catalytic activity of 6PGD. Mutagenesis studies, where specific amino acids within the active site are modified or substituted, have provided insights into the role of these residues in catalysis.

4. Getting Ribose-5-phosphate

Ribose-5-phosphate is the desired product of the reaction. It is a five-carbon sugar-phosphate molecule that serves as a precursor for the synthesis of nucleotides (building blocks of DNA and RNA) and coenzymes (such as NADH, NADPH, FAD, and B12).  The pentose phosphate pathway is a metabolic pathway that operates alongside glycolysis to generate important molecules such as ribose-5-phosphate (ribose-5-P) and reducing equivalents in the form of NADPH. Ribose-5-phosphate isomerase interconverts these two phosphorylated sugars via an enediol intermediate.



An aldose is converted into a ketose through an isomerization reaction. The conversion of ribulose-5-P to ribose-5-P involves several steps:

The conversion of ribulose-5-phosphate to ribose-5-phosphate in the pentose phosphate pathway involves the following steps:

Isomerization: Ribulose-5-phosphate is converted to ribose-5-phosphate through the action of ribulose-5-phosphate epimerase. This enzyme catalyzes the rearrangement of the hydroxyl groups at the C1 and C2 carbons of the ribulose-5-phosphate molecule, resulting in the formation of ribose-5-phosphate.

Oxidative Decarboxylation: Ribose-5-phosphate is then acted upon by ribulose-5-phosphate isomerase. This enzyme catalyzes the oxidative decarboxylation of ribose-5-phosphate, leading to the formation of ribulose-5-phosphate. This step involves the removal of a carbon dioxide (CO2) molecule and the reduction of NADP+ to NADPH.

Phosphorylation: Finally, ribulose-5-phosphate is converted back to ribose-5-phosphate through the action of phosphopentose isomerase. This enzyme catalyzes the rearrangement of the phosphate group on the C1 carbon of ribulose-5-phosphate, resulting in the formation of ribose-5-phosphate.

These enzymatic reactions ensure the interconversion of ribulose-5-phosphate and ribose-5-phosphate in the pentose phosphate pathway, allowing for the synthesis of important molecules such as nucleotides and coenzymes.
The net result of these steps is the conversion of ribulose-5-P to ribose-5-P, an essential intermediate in various biosynthetic pathways. Ribose-5-P is utilized in the biosynthesis of coenzymes, including NADH, NADPH, FAD, and B12, which play crucial roles in cellular metabolism. Additionally, ribose-5-P serves as a precursor for the synthesis of nucleotides and nucleic acids, such as DNA and RNA, which are vital for cellular processes like DNA replication and protein synthesis. This is a crucial step in the pentose phosphate pathway and is essential for life. Ribose-5-phosphate is a key intermediate that serves as a precursor for the synthesis of nucleotides and coenzymes, which are vital for various cellular processes.

Besides being the building blocks of DNA and RNA, the genetic materials that store and transmit genetic information nucleotides are also essential for the synthesis of ATP (adenosine triphosphate), the energy currency of the cell. Ribose-5-phosphate provides the necessary carbon and energy source for nucleotide synthesis. Coenzymes, such as NADH, NADPH, FAD, and B12, play also critical roles in cellular metabolism. NADH and NADPH are involved in redox reactions, serving as electron carriers that participate in energy-producing reactions (such as cellular respiration) and biosynthetic pathways (such as fatty acid and cholesterol synthesis). FAD is a cofactor for enzymes involved in energy production, such as the citric acid cycle. Vitamin B12 (cobalamin) is a coenzyme required for various enzymatic reactions, including those involved in nucleotide and amino acid metabolism. Therefore, the production of ribose-5-phosphate from ribulose-5-P is essential for the synthesis of nucleotides, coenzymes, and other important biomolecules. These molecules are involved in crucial cellular processes, including DNA replication, RNA synthesis, energy production, and the maintenance of cellular redox balance. Without the pentose phosphate pathway and the conversion of ribulose-5-P to ribose-5-P, the cell would lack the necessary building blocks and coenzymes required for these vital processes, which would severely impair its ability to function and survive.

In the second stage of the pentose phosphate pathway, ribulose-5-phosphate (Ru5P) undergoes isomerization and epimerization reactions catalyzed by ribulose-5-phosphate isomerase and ribulose-5-phosphate epimerase, respectively. Ribulose-5-phosphate isomerase catalyzes the conversion of Ru5P to ribose-5-phosphate (R5P). This isomerization reaction involves rearrangement of the carbon skeleton, resulting in the formation of R5P. R5P is an important precursor in the biosynthesis of nucleotides, which are essential for DNA and RNA synthesis. Therefore, in rapidly dividing cells with increased DNA synthesis, the production of R5P is relatively high. On the other hand, ribulose-5-phosphate epimerase catalyzes the conversion of Ru5P to xylulose-5-phosphate (Xu5P). This epimerization reaction involves the exchange of specific functional groups on the molecule, resulting in the formation of Xu5P. The relative amounts of R5P and Xu5P produced from Ru5P depend on the metabolic needs of the cell. If the pentose phosphate pathway is primarily utilized for NADPH production, the ratio of Xu5P to R5P is approximately 2:1. Xu5P and R5P can then be further metabolized into glycolytic intermediates in the third stage of the pathway. The regulation of R5P and Xu5P production is tightly controlled to meet the demands of the cell. The pathway can be upregulated or downregulated depending on the metabolic requirements. Rapidly dividing cells or cells in need of nucleotide biosynthesis will favor the production of R5P, while cells requiring NADPH production will produce Xu5P in larger amounts. These isomerization and epimerization reactions, similar to the reaction catalyzed by triose phosphate isomerase, are thought to occur via enediolate intermediates. Enediolate intermediates are highly reactive species that play a crucial role in the rearrangement of functional groups and carbon skeleton during these reactions.

Right-handed chiral form of Ribose-5-phosphate

Ribose-5-phosphate is a five-carbon sugar molecule that is right-handed chiral. Chirality refers to the property of an object that cannot be superimposed onto its mirror image. In the case of ribose-5-phosphate, its chiral nature is determined by the arrangement of atoms around its asymmetric carbon atom, also known as the chiral center. The chiral center in ribose-5-phosphate is the carbon atom located in the second position of the sugar ring. It is called the C2 carbon. This carbon is attached to four different chemical groups or functional groups: a hydroxyl group (-OH), a phosphate group (-PO₄), a hydrogen atom (-H), and the rest of the sugar ring structure. The presence of these distinct groups creates two possible configurations around the chiral center: the D-configuration (right-handed) and the L-configuration (left-handed). In the case of ribose-5-phosphate, it adopts the D-configuration, making it right-handed. The specific arrangement of these groups around the chiral center is determined during the biosynthesis of ribose-5-phosphate. Enzymes and cellular processes control the synthesis of this molecule, ensuring the correct arrangement of atoms and groups to achieve the D-configuration. It's important to note that chirality is a fundamental property of many biological molecules, including sugars, amino acids, and nucleic acids. The chiral nature of these molecules plays a crucial role in their interactions with other molecules and their biological functions. The right-handed chirality of ribose-5-phosphate is determined by enzymes involved in its biosynthesis. Specifically, the enzyme ribose-5-phosphate isomerase is responsible for interconverting ribulose-5-phosphate and ribose-5-phosphate. This enzyme plays a crucial role in establishing the chiral configuration of ribose-5-phosphate. Ribose-5-phosphate isomerase catalyzes the isomerization reaction, which involves rearranging the double bond in ribulose-5-phosphate to form an enediol intermediate. This intermediate then undergoes tautomerization and rearrangement to ultimately yield ribose-5-phosphate. The specific mechanism by which ribose-5-phosphate isomerase achieves right-handed chirality is not fully understood. It is likely that the enzyme's active site and binding interactions with the substrate play a significant role in determining the stereochemistry of the product. While the right-handed chirality of ribose-5-phosphate is essential for its biological function, there is no inherent physical necessity for it to be in the D-configuration. In theory, it could exist in a mixed or racemic form, or even in the L-configuration. However, in the context of biological systems, the specific D-configuration of ribose-5-phosphate is critical for its involvement in nucleotide and nucleic acid synthesis. The right-handed chirality of ribose-5-phosphate is crucial for proper polymerization and the formation of biologically functional nucleotides and nucleic acids, such as DNA and RNA. The D-configuration of ribose-5-phosphate allows for the correct alignment and bonding with other components, ensuring the structural integrity and stability of these macromolecules.

If ribose were racemic or mixed in its chiral configuration, it would have significant implications for the formation and function of nucleotides, which are crucial for life as we know it. The chiral nature of ribose, specifically its right-handed configuration (D-ribose), is essential for the proper assembly and stability of nucleotides and nucleic acids, such as DNA and RNA. Nucleotides are composed of three components: a sugar molecule (ribose or deoxyribose in the case of DNA), a phosphate group, and a nitrogenous base. The sugar molecule, ribose, plays a vital role in providing the structural framework for nucleotide polymerization. In nucleotide polymerization, the sugar molecules (ribose) link together through a phosphodiester bond formation, creating a backbone for the nucleic acid chain. The specific arrangement of the sugar molecules in the backbone is critical for the stability and fidelity of the nucleic acid structure. If ribose were racemic or mixed in its chiral configuration, it would result in the incorporation of both right-handed (D) and left-handed (L) sugar molecules into the nucleotide chain. This mixed configuration would introduce structural irregularities and disrupt the stability and integrity of the nucleic acid structure. The polymerization process would be hindered, leading to a loss of the specific information-carrying capacity and functionality of nucleic acids. Furthermore, the recognition and binding of nucleotides with other molecules, such as enzymes and proteins, rely on the specific shape and arrangement of the sugar molecules. A racemic or mixed configuration of ribose would hinder proper binding interactions and compromise the overall functionality of nucleotides. Therefore, the right-handed chirality of ribose, which is achieved through the precise arrangement of atoms and the involvement of enzymes during biosynthesis, is crucial for the formation, stability, and functionality of nucleotides. The specific design of ribose as a right-handed chiral molecule point to the necessity for life's fundamental processes, including the storage and transmission of genetic information.

While there is no physical necessity for ribose-5-phosphate to be in the D-configuration, the fact that it adopts this configuration highlights the precision and foresight involved in its design. This intentional arrangement ensures that ribose-5-phosphate can properly interact and bond with other molecules in the cell, particularly in the context of nucleotide and nucleic acid synthesis. The existence of intricate and purposeful molecular designs throughout living systems suggests that an intelligent agent carefully designed these systems to fulfill specific functions. The precise arrangement of molecules like ribose-5-phosphate points to an intentional design that enables the formation of nucleotides, which are essential for the storage and transmission of genetic information.

Further Essential Structural Features of Ribose for Biological Function

In addition to being right-handed chiral, several other structural features are essential for ribose to perform its function in biological processes:  Ribose contains hydroxyl (-OH) groups attached to its carbon atoms. These hydroxyl groups play a crucial role in forming bonds with other molecules during the synthesis of nucleotides and nucleic acids. They are involved in phosphodiester bond formation, where the hydroxyl group of one ribose molecule reacts with the phosphate group of another molecule, creating a backbone for nucleotide polymerization.  Ribose is a pentose sugar, meaning it has a five-carbon backbone. This pentose structure provides the necessary flexibility and rigidity to form stable and functional nucleotide chains. The specific arrangement of carbon atoms in the pentose backbone ensures the proper alignment of functional groups and allows for efficient bonding with other components.
If ribose were not a pentose sugar and instead had a different carbon backbone, it would significantly impact the structure and function of nucleotides and nucleic acids.  The specific arrangement of five carbon atoms allows for efficient bonding with nitrogenous bases and phosphate groups. If ribose had a different carbon backbone, it might result in an unstable or less rigid structure, making the nucleotide chains more susceptible to breakage or distortion. Enzymes and proteins involved in nucleotide synthesis, DNA replication, and RNA transcription are specifically adapted to interact with the pentose backbone of ribose. A different carbon backbone would likely be incompatible with these enzymes and proteins, hindering their ability to recognize and interact with ribose during crucial cellular processes. There is interdependence between the specific structure of ribose (including its pentose backbone) and the enzymes and proteins involved in nucleotide synthesis, DNA replication, and RNA transcription. The compatibility between ribose and these biomolecules is essential for their proper functioning in cellular processes. Enzymes are highly specific in their recognition and interaction with substrates. They have active sites that are precisely shaped to accommodate specific molecules and facilitate chemical reactions. In the case of enzymes involved in nucleotide synthesis, DNA replication, and RNA transcription, their active sites are adapted to interact with the pentose backbone of ribose. The pentose backbone provides a specific arrangement of carbon atoms and functional groups that enzymes and proteins recognize and bind to. The enzymes involved in these processes have to interact with ribose and its specific structure, allowing for efficient catalysis and proper incorporation of ribose into nucleotide chains. If the carbon backbone of ribose were different, enzymes and proteins would not be able to recognize and interact with ribose effectively. This would hinder their ability to catalyze the necessary reactions for nucleotide synthesis, DNA replication, and RNA transcription. As a result, the overall processes required for the maintenance and transmission of genetic information would be compromised. The interdependence between ribose structure and enzyme/protein recognition highlights the intricate relationship between biomolecules in biological systems. It showcases the specificity and precision required for the successful execution of crucial cellular processes.

The interdependence between the specific structure of ribose and the enzymes and proteins involved in nucleotide synthesis, DNA replication, and RNA transcription are clear evidence of intelligent design and a strong indication that these components were created together, and fit to interact harmoniously from the start. The intricate compatibility between ribose and the enzymes and proteins involved in these processes suggests a purposeful arrangement. The precise shape of enzyme active sites and their ability to recognize and interact specifically with the pentose backbone of ribose indicate a well-designed system. This implies that the functionality and interplay of ribose, nucleic acids (RNA and DNA), and the associated enzymes were intentionally orchestrated to work together seamlessly. A stepwise origin, where each component gradually develops independently, would be highly unlikely to result in the intricate interdependence observed in these cellular processes. The specificity and precision required for the recognition, binding, and catalytic activities of enzymes towards ribose strongly suggest that they were co-designed with ribose and nucleic acids, rather than being assembled in a gradual, incremental manner. The intricate interdependence between ribose, nucleic acids, and the enzymes involved in their synthesis and replication highlights the concept of irreducible complexity. This concept posits that certain biological systems require the simultaneous presence and precise coordination of multiple components to function properly. The interdependent relationship between ribose, nucleic acids, and enzymes aligns with this concept, as any alteration or absence of these components would hinder the essential processes involved in genetic information storage and transmission.

The specific arrangement of carbon atoms in ribose's pentose structure facilitates proper base pairing between nucleotides. In DNA, adenine (A) pairs with thymine (T), and guanine (G) pairs with cytosine (C). In RNA, uracil (U) replaces thymine. If the carbon backbone were different, the complementary base pairing might be disrupted or altered, leading to the incorrect encoding and transmission of genetic information. The pentose structure of ribose plays a vital role in nucleotide polymerization. During DNA and RNA synthesis, the phosphate group of one nucleotide forms a phosphodiester bond with the hydroxyl group of the pentose sugar in the adjacent nucleotide, creating a stable polymer chain. A different carbon backbone could hinder the formation of these bonds, affecting the polymerization process and compromising the integrity of nucleic acids. Ribose-5-phosphate, the phosphorylated form of ribose, plays a critical role in cellular metabolism. The phosphate group (-PO₄) attached to ribose provides the necessary energy for various biochemical reactions and is essential for the synthesis of nucleotides and other cellular components. Ribose can form glycosidic bonds with nitrogenous bases, such as adenine, guanine, cytosine, and uracil (in RNA) or thymine (in DNA). These bonds are crucial for the formation of nucleosides and nucleotides, where the nitrogenous base is attached to the ribose sugar. The specific arrangement and bonding between ribose and the nitrogenous base determine the genetic information encoded in DNA or RNA.  The stability of ribose is vital for the integrity and functionality of nucleic acids. Ribose undergoes various chemical modifications and interactions within the cellular environment to maintain its structural integrity and protect it from degradation. Enzymes and cellular processes regulate these modifications and ensure the stability of ribose during nucleotide synthesis and nucleic acid metabolism.

Gaspar Banfalv Ribose Selected as Precursor to Life (2019): The critical chemical reactions yielding the first informational macromolecule have yet to be clarified.

Molecular modeling studies have revealed that of the four pentoses tested (ribose, arabinose, xylose, and lyxose), only β-D-ribose could be successfully inserted into nucleotides while maintaining the necessary free rotation for functional groups (hydroxyl, phosphate, and base) and overall stability.  Unrestricted rotation of functional groups in ribonucleotides is crucial for their structural flexibility and ability to form double-stranded structures. The selection of β-D-ribose allows for the necessary spatial arrangement and freedom of rotation, enabling proper nucleotide function.  Attempting to fit any other pentose sugar into nucleotides creates a steric hindrance or barrier. This suggests that alternative pentose sugars, such as arabinose, xylose, or lyxose, would not be compatible with the necessary functional group arrangement and would hinder the formation of functional ribonucleic acids. The hypothetical lyxonucleotides, derived from lyxose, face the issue of functional groups being within van der Waals radius distance, preventing their free rotation. This likely explains why lyxonucleotides do not exist in nature. Selecting β-D-ribose over α-D-ribose provides an advantage in terms of the spatial arrangement and localization of functional groups. The positioning of the C1'-base and C2'-OH substituents of ribose in nucleotides allows them to be distantly located, promoting freedom of movement and flexibility.

Ribose, the sugar component of RNA is not readily available as a building block for prebiotic RNA synthesis. This raises questions about the feasibility of RNA formation under abiotic conditions. Chemical obstacles are arguments against the spontaneous formation of nucleic acids, including RNA, under abiotic conditions. The ester linkage between ribose and phosphoric acid, which forms the backbone of RNA, is prone to hydrolysis. This raises concerns about the stability and persistence of RNA molecules over long periods of time. The attachment of nucleobases to the ribose-phosphate backbone to form a random nucleotide sequence was followed by the hydrolysis of the polymer chain. This hydrolysis can break down the RNA molecule and limit its ability to persist and function.

While the light elements (CHNOPS) are crucial for biological processes,hydrogen and helium, the lightest elements, are more abundant in the Universe but less prevalent on Earth due to their loss through processes like Planetary Air Leak. This scarcity of light elements on Earth could pose a challenge to the spontaneous generation and accumulation of the necessary elements for the development of complex biological systems.  There is also the scarcity of ribose as a building block for prebiotic RNA. Ribose is crucial for the formation of the ribose-phosphate backbone in RNA, and its scarcity would hinder the spontaneous synthesis of prebiotic RNA molecules. This scarcity raises questions about how the necessary ribose molecules would have been available in sufficient quantities to support the formation of genetic information. The ester linkage between ribose and phosphoric acid is prone to hydrolysis, which can degrade RNA molecules. This chemical obstacle poses a challenge to the stability and preservation of genetic information over extended periods of time. It raises questions about how the necessary mechanisms for protecting and preserving genetic information, such as enzymes and repair systems, could have emerged through natural processes.  Only β-D-ribose can be effectively incorporated into nucleotides due to its compatibility with the functional groups (OH, phosphate, and base). This implies a specific configuration and interaction between ribose and other components of nucleotides. The presence of a sterical barrier when trying to fit other pentose sugars into nucleotides raises questions about how the precise compatibility and specificity between the components of genetic molecules could have arisen through random, unguided processes. These potential problems and challenges, from an intelligent design perspective, suggest that the intricate compatibility and interdependence of the bioelements and chemical structures involved in genetic information raise questions about the plausibility of a stepwise, gradual origin of such complex systems. They prompt consideration of alternative explanations that involve purposeful design or guidance in the development of these essential biological components.

Ribulose-5-phosphate epimerase

Ribulose-5-phosphate epimerase (RPE) is an enzyme involved in carbohydrate metabolism. It catalyzes the interconversion of ribulose-5-phosphate (R5P) and xylulose-5-phosphate (X5P), which are important intermediates in the pentose phosphate pathway and glycolysis. Ribulose-5-phosphate epimerase can exist as both monomeric and dimeric forms, depending on the organism and specific conditions. In some organisms, such as Escherichia coli (E. coli), the enzyme is monomeric. The primary function of RPE is to catalyze the reversible epimerization reaction between R5P and X5P. This conversion involves the rearrangement of functional groups within the sugar molecule.
The average size of RPE varies among different organisms, but it typically consists of around 200-300 amino acids. The enzyme has a total structure weight of 153.58 kDa and contains 11,048 atoms in E.Coli.
The exact mechanism of the RPE-catalyzed reaction involves the transfer of a hydride ion between carbon atoms, leading to the conversion of R5P to X5P or vice versa. The precise details of the mechanism may vary depending on the specific enzyme and organism. The mechanism of this reaction involves several steps. R5P initially binds to the active site of the enzyme. The active site of RPE contains amino acid residues that interact with the substrate and stabilize its binding. The first step in the mechanism is the isomerization of R5P. The enzyme facilitates the rearrangement of the carbon backbone, leading to the formation of an intermediate. The key step in the mechanism involves the transfer of a hydride ion (H-) between carbon atoms. This hydride transfer leads to the conversion of R5P to X5P or vice versa. The enzyme provides an appropriate environment within its active site to facilitate this transfer and maintain the stability of the reaction. Following the hydride transfer, the converted product, either X5P or R5P, is released from the active site, making it available for further metabolic pathways. The three-dimensional structure of the enzyme and the arrangement of amino acids in its active site play a critical role in enabling the precise hydride transfer reaction.



The origin of an enzyme like Ribulose-5-phosphate epimerase (RPE) can be seen as the result of purposeful design and engineering. Enzymes are remarkable molecular machines that exhibit incredible precision, efficiency, and specificity in catalyzing biochemical reactions. The precise arrangement and fine-tuning of amino acids in the active site of RPE, as well as the overall three-dimensional structure of the enzyme, are essential for its specific recognition and binding of substrates. This precision allows for the highly specific catalysis of the hydride transfer reaction between R5P and X5P. The intricate coordination of charges, shapes, and other molecular features within the active site of RPE, which enable its catalytic activity, cannot be adequately explained by random chance or unguided natural processes. The level of complexity and optimization observed in enzymes suggests the involvement of an intelligent agent, capable of designing and fine-tuning these molecular systems to perform specific functions. Additionally, the fact that RPE exhibits high specificity and selectivity in recognizing and binding its substrates further supports the notion of design. The enzyme's active site possesses the necessary amino acid residues and atoms to interact with the substrate and stabilize its binding, facilitating the precise hydride transfer reaction. Such a level of specificity and functionality implies intentional design rather than a gradual and random evolutionary process.

Ribulose-5-phosphate isomerase

Ribulose-5-phosphate isomerase (RPI), also known as ribose-5-phosphate isomerase,  the reversible isomerization of ribulose-5-phosphate (R5P) to ribose-5-phosphate (R5P) in the pentose phosphate pathway. This enzymatic reaction plays a crucial role in the interconversion of sugars and the generation of important metabolic intermediates. The average size of RPI varies among different organisms, but it typically consists of around 200-300 amino acids. The enzyme has a total structure weight of 96.42 kDa and contains 6,964 atoms in E.Coli. The enzyme's structure may contain one or more metal cofactors, such as magnesium or manganese, in its active site. These metal ions are essential for catalytic activity and are often coordinated by specific amino acid residues within the enzyme. The metal cofactor is coordinated by specific amino acid residues within the active site of RPI. These residues act as ligands, binding to the metal ion and forming coordination bonds. The coordination of the metal ion by the amino acid residues helps stabilize the ion and enables its participation in the catalytic reaction. The specific amino acid residues involved in metal coordination can vary among different RPI enzymes. However, common amino acids that often participate in metal ion binding include histidine (His), aspartate (Asp), and glutamate (Glu). These amino acids have side chains that contain functional groups capable of coordinating with metal ions through coordination bonds. The presence of the metal cofactor in the active site enhances the enzyme's catalytic activity by providing additional coordination sites and facilitating the proper positioning of the substrate for the isomerization reaction. The metal ion can interact with the functional groups of the substrate, promoting the necessary rearrangements and stabilizing reaction intermediates. The specific coordination geometry and interactions between the metal cofactor and the amino acid residues in the active site are crucial for the enzyme's function. The intricate coordination of charges, shape, and other molecular features within the active site is essential for the specific recognition and binding of the metal ion and the substrate. Any deviation from the precise arrangement may impair the enzyme's catalytic activity. In terms of synthesis, the metal cofactors required by RPI are typically obtained from the cellular environment. Organisms have specific mechanisms for acquiring and regulating the levels of essential metal ions such as magnesium and manganese. These metal ions can be obtained through uptake from the environment or through transport systems within the cell. Once inside the cell, the metal ions are likely incorporated into the active site of RPI through protein-metal interactions. Regarding the origin of the fine-tuning of metal coordination in enzymes like RPI, the precise arrangement of amino acid residues and metal ions in the active site points to the involvement of an intelligent agent in the design and creation of these systems. The intricate coordination and optimal positioning of metal cofactors and substrate molecules within the active site require a level of specificity and complexity that is unlikely to arise through natural, unguided processes. 

RPI functions by binding the substrate, R5P, in its active site and facilitating the rearrangement of chemical bonds. The precise arrangement of amino acids within the active site is crucial for substrate recognition and binding. Specific amino acid residues, including histidine, glutamate, and lysine, are involved in coordinating the metal cofactor and forming interactions with the substrate. These interactions stabilize the transition state and promote the isomerization reaction. The substrate specificity of RPI is highly specific to ribulose-5-phosphate (R5P). The active site of the enzyme recognizes the specific shape and functional groups of R5P, allowing for precise binding and subsequent isomerization. The active site typically consists of several binding sites to accommodate the substrate and facilitate the catalytic reaction. The regulation of RPI can occur through various mechanisms, including allosteric regulation and post-translational modifications. Allosteric effectors or specific metabolites may regulate the enzyme's activity, ensuring its participation in metabolic pathways based on the cellular needs. Post-translational modifications, such as phosphorylation or acetylation, can also modulate RPI activity. Regarding atoms, metal cofactors, such as magnesium or manganese ions, are crucial for the catalytic activity of RPI. These metal ions are coordinated by specific amino acid residues within the enzyme's active site, facilitating the isomerization reaction. The precise arrangement of amino acids and the coordination of charges and shape within the active site are essential for specific substrate recognition and binding. The precise rotation angles of atoms, including those of certain amino acids, can be crucial for the catalytic activity of enzymes like RPI. Optimal rotation angles allow for the proper alignment and interaction of functional groups within the active site, enabling the catalytic event. Deviations in these angles may disrupt the necessary molecular interactions and compromise enzymatic activity. The origin of the fine-tuning of rotation angles in enzymes can be attributed to Intelligent Design. An intelligent designer can purposefully engineer the precise arrangement of atoms, amino acids, and their rotation angles to maximize catalytic efficiency and achieve specific functions.

Phosphopentose isomerase

Phosphopentose isomerase is responsible for the production of ribose-5-phosphate, a key molecule used in nucleotide synthesis and other cellular processes. Phosphopentose isomerase catalyzes the isomerization of ribulose-5-phosphate to ribose-5-phosphate. This is a reversible reaction that involves rearranging the carbon backbone of the sugar molecule. The conversion of ribulose-5-phosphate to ribose-5-phosphate is an important step in the pentose phosphate pathway, as ribose-5-phosphate serves as a precursor for the synthesis of nucleotides and other important biomolecules. Phosphopentose isomerase is typically composed of a single polypeptide chain, making it a monomeric enzyme. It is primarily made up of amino acids arranged in a specific sequence determined by the gene encoding the enzyme. The three-dimensional structure of phosphopentose isomerase allows it to interact with its substrate and catalyze the isomerization reaction efficiently. Phosphopentose isomerase specifically acts on ribulose-5-phosphate as its substrate. The enzyme recognizes and binds to the substrate through specific interactions between the active site of the enzyme and the substrate molecule. The binding site within the enzyme facilitates the conversion of ribulose-5-phosphate to ribose-5-phosphate.  Phosphopentose isomerase is found in various organisms, including bacteria, plants, and animals. It plays a crucial role in cellular metabolism by providing ribose-5-phosphate for nucleotide synthesis and other biochemical pathways. The enzyme is often conserved across different species, indicating its importance in biological processes.

The DNA structure

The central dogma of molecular biology states that genetic information flows from DNA to RNA to proteins, and this process plays a fundamental role in determining the shapes and activities of individual cells. The genetic instructions encoded in DNA sequences are transcribed into RNA molecules, which are then translated into specific amino acid sequences to form proteins. Proteins are essential components of cells and perform various functions. Many proteins are enzymes that catalyze biochemical reactions involved in cellular metabolism. Enzymes facilitate and regulate the chemical reactions required for processes such as energy production, nutrient metabolism, and synthesis of biomolecules. These metabolic processes are vital for cell survival, growth, and function. In addition to enzymatic functions, proteins also have structural roles, providing support and organization to cellular components. Structural proteins contribute to the formation of cell membranes, cytoskeleton, and extracellular matrix, maintaining cell shape and integrity. They are involved in cell adhesion, cellular movement, and the maintenance of tissue architecture. Furthermore, proteins play regulatory roles in cells. They can act as switches, turning on or off specific cellular processes in response to signals or changes in the cell's environment. Regulatory proteins are involved in gene expression, signal transduction, and cell signaling pathways, allowing cells to respond and adapt to internal and external cues. Some proteins are directly involved in maintaining and transmitting genetic information. For example, DNA-binding proteins and histones help package and organize DNA into chromatin structures, allowing proper DNA replication, transcription, and gene regulation. RNA-binding proteins interact with RNA molecules to control their stability, localization, and translation, influencing gene expression. The central dogma highlights the importance of DNA, RNA, and proteins in the intricate workings of cells. Genetic information encoded in DNA ultimately determines the sequence of amino acids in proteins, which in turn dictate the structure and function of cells. This interplay between DNA, RNA, and proteins is fundamental to the understanding of molecular biology and the complex processes underlying cellular life.

This interplay between DNA, RNA, and proteins is evidence of a complex and intricately designed system. The interdependence and coordinated functioning of these components suggest a purposefully designed implementation rather than originating from a gradual unguided process. The complex and integrated nature of cellular processes indicates the involvement of a cognitive agent or a goal-directed process. The powerful molecular control networks guarantee the functional coherence of the cell and the invariance of its basic chemical scheme. This coherence and integrated complexity are confirmatory evidence for intelligent design. The intricate molecular machinery involved in processes such as transcription and translation is a hallmark of intelligent design. These tiny, intricately constructed molecular machines, arranged in cooperative systems, provide the cell with the necessary functionality. The simultaneous emergence of these components and their precise molecular interactions indicate a purposeful and carefully planned invention rather than a step-by-step process. The chemical architecture of life and its crucial reactions could not have emerged one after the other by chance. A series of exquisitely designed biomolecules and molecular machines had to be in place at the same time to make life's molecular workforce, proteomes, function properly. This coordinated complexity, characterized by exquisite design details down to the atomic level, is evidence of foresight, sound engineering, and high technology. The interplay between DNA, RNA, and proteins is a highly intricate network that is crucial for life. The interplay and balance among these components indicate the planning and engineering marvel involved in their design.

DNA and RNA, the two types of nucleic acids, play crucial roles in storing and transmitting genetic information within cells. The structures of these molecules reflect several key principles: Genetic information must be stored in a form that is compact and stable over long periods. Both DNA and RNA achieve this by utilizing the double helix structure, where two complementary strands are held together by hydrogen bonds. This structure allows for efficient storage of information while maintaining stability. Genetic information stored in DNA needs to be decoded and utilized by the cell. Transcription is the process by which the genetic sequence of DNA is copied onto RNA molecules, enabling them to direct protein synthesis during translation. This process ensures that the information in DNA is accessible and usable by the cellular machinery.  Information contained in DNA or RNA needs to be accessible to proteins and other nucleic acids. Various cellular components, including proteins, must recognize specific nucleic acid sequences and bind to them in a manner that modulates their function. These sequence-specific interactions allow for the regulation of gene expression, DNA replication, and other essential cellular processes.  Genetic information must be faithfully passed on to the progeny. DNA achieves this through replication, where an exact copy of the DNA molecule is produced, ensuring that each daughter cell receives the same set of instructions as the parent. This replication process is crucial for maintaining the continuity of genetic information across generations. Nucleic acids, particularly RNA, are not merely inert "read-only" molecules. RNA, with its single-stranded nature, exhibits dynamic properties and serves additional roles beyond information storage. It can act as a structural scaffold and display catalytic activity in processes that decode genetic information. These properties allow RNA to contribute to diverse cellular functions. Understanding the structural properties of nucleic acids, their interactions with proteins, and their dynamic nature is essential for comprehending how they carry out their vital functions within cells. By unraveling these intricacies, we gain insights into the fundamental mechanisms that underlie the storage, transmission, and utilization of genetic information in living organisms.

The ATGC quartet, consisting of adenine, thymine, guanine, and cytosine, provides the foundation for the chemical architecture of DNA. The specific arrangement and pairing of these bases allow for the compact storage of genetic information in a stable manner. This finely engineered design exhibits exquisite balance and interplay, serving as a hallmark of foresight and sound engineering.  The replication of DNA, where an exact copy is made to pass on genetic instructions, requires a highly intricate network of molecular machinery. The process involves the coordination of multiple enzymes, proteins, and regulatory factors, all working in harmony to ensure the faithful transmission of information. Such a complex system indicates the involvement of ingenious solutions and carefully planned inventions. Transcription of DNA into RNA and the subsequent translation of RNA into proteins involve a series of remarkable biochemical reactions and interactions. The precise recognition of nucleic acid sequences by proteins and other molecules, along with their specific binding, showcases the incredible bioengineering behind these processes. This level of mastery and engineering cleverness suggests a purposeful design rather than a product of blind chemical forces. Furthermore, the dynamic properties of RNA, its ability to serve as a structural scaffold, and its catalytic proficiency in decoding genetic information add to the wonder of its design. These features demonstrate the intricate interplay and exquisitely engineered molecular arrangement necessary for RNA to contribute to the functioning of cells. The origin of these complex and interdependent features, such as information storage capacity, specific recognition, faithful replication, and dynamic functionality, cannot be explained by gradual naturalistic - evolutionary processes. The simultaneous emergence of these features, their finely tuned interactions, and the incredible bioengineering required all point towards the involvement of a super-intelligent designer.

DNA, the genetic material found in living organisms, exists in various structural forms depending on its environment and base sequence. While the most common form is known as B-DNA, which was initially described by James Watson, Francis Crick, Rosalind Franklin, and others, there are other conformations worth mentioning. B-DNA, the canonical form, exhibits several key structural features. It consists of two polynucleotide strands that run in opposite directions (antiparallel) and coil around a central axis, forming a right-handed double helix with a diameter of approximately 20 angstroms. The nucleotide bases, which include adenine (A), thymine (T), guanine (G), and cytosine (C), pair through hydrogen bonds and are almost perpendicular to the helix axis. In B-DNA, the bases are nestled inside the helix, while the sugar-phosphate backbones wind along the outside, creating major and minor grooves. The base pairs are exposed only at the edges to the surrounding solvent. The symmetry of the DNA molecule remains intact regardless of the specific base composition, with each base pair maintaining a similar width. This property allows base pairs like A-T and G-C to interchange positions without altering the structure, while other combinations would significantly distort the helix. The ideal B-DNA helix has 10 base pairs per turn, resulting in a helical twist of 36 degrees per base pair, and a pitch (rise per turn) of 34 angstroms.

However, DNA can adopt alternative conformations in different solvent conditions or with specific base sequences. One such conformation is Z-DNA, which forms a left-handed helix instead of the right-handed helix of B-DNA. Z-DNA is characterized by a zigzag backbone, arising from a different arrangement of the sugar-phosphate backbone. It occurs under high salt concentrations or with specific DNA sequences that contain alternating purine-pyrimidine repeats. Another notable DNA structure is A-DNA, which resembles B-DNA but exhibits a shorter and wider helix with a pitch of 28 angstroms and 11 base pairs per turn. A-DNA can form under dehydrated conditions or with specific DNA sequences that have a high DNA strand concentration. It is often transient and not as prevalent as B-DNA. Moreover, DNA can exhibit variations in structure and flexibility due to factors such as supercoiling, protein binding, and DNA damage. These alterations play crucial roles in processes like DNA replication, transcription, and DNA repair.

To achieve the coiling of DNA into a double helix structure, several factors are necessary. These factors are essential for the stability and functionality of DNA, which is crucial for life:  The complementary base pairing between adenine (A) and thymine (T), as well as guanine (G) and cytosine (C), forms the foundation of DNA's double helical structure. The hydrogen bonds between these base pairs contribute to the stability of the helix. The phosphodiester bonds connect the sugar moieties of adjacent nucleotides within each DNA strand. These bonds provide the backbone for the DNA molecule and contribute to its structural integrity. The two polynucleotide strands in DNA run in opposite directions, with one strand oriented in the 5' to 3' direction and the other in the 3' to 5' direction. This antiparallel arrangement is crucial for the proper base pairing and allows for the formation of hydrogen bonds between the bases.  The sugar-phosphate backbone of DNA provides structural support and protection for the nitrogenous bases. The sugar molecules (deoxyribose) and phosphate groups alternate along the length of the DNA strands, forming the external framework of the double helix.  The hydrogen bonds between the complementary base pairs (A-T and G-C) hold the two DNA strands together. These bonds are relatively weak individually but collectively contribute to the overall stability of the double helix.

The coiling of DNA into a double helix is essential for life due to the following reasons: DNA carries the genetic information necessary for the functioning and development of all living organisms. The double helical structure allows DNA to store this information in a compact and stable manner. The sequence of the bases along the DNA strands encodes the instructions for protein synthesis and other cellular processes.  During DNA replication, the double helix unwinds, and each strand serves as a template for the synthesis of a new complementary strand. The ability of DNA to coil into a stable structure ensures accurate replication, allowing for the faithful transmission of genetic information from one generation to the next.  The double helical structure of DNA plays a crucial role in gene expression. Various proteins, such as transcription factors and RNA polymerases, bind to specific regions of DNA to regulate gene expression. The accessibility of these regulatory regions is influenced by the coiling and structural organization of DNA.  In eukaryotic cells, DNA is packaged into chromosomes to fit within the nucleus. The coiling of DNA into higher-order structures, such as nucleosomes and chromatin, enables efficient packaging while still allowing access to the genetic information when needed. The double helical structure provides stability to the DNA molecule, protecting it from chemical and physical damage. It helps shield the nitrogenous bases within the helix, reducing their exposure to potentially harmful agents. Additionally, the coiled structure of DNA helps to prevent DNA strands from tangling or forming knots. DNA carries an incredibly complex and specified genetic code. The sequence of nucleotide bases along the DNA strands encodes the instructions for the development, functioning, and regulation of all living organisms. The precise arrangement of the bases, their pairing rules, and the complementary nature of the strands require highly specific and specified information. The generation of such information through random, unguided processes is astronomically improbable. Intelligent agency, however, is known to produce complex and specified information, as seen in human-designed systems.  DNA replication is a highly accurate process that ensures the faithful transmission of genetic information from one generation to the next. The ability of DNA to unwind, separate its strands, and serve as templates for the synthesis of new complementary strands is a remarkably precise and coordinated mechanism. Random processes, devoid of guidance, are unlikely to produce the intricate molecular machinery required for accurate DNA replication.   The double helical structure of DNA plays a crucial role in gene expression regulation. It provides a platform for the binding of various proteins, such as transcription factors and RNA polymerases, which control the activation or repression of specific genes. The specific positioning and accessibility of regulatory regions within the DNA molecule require precise structural organization. The existence of finely tuned gene regulatory networks implies the need for intentional design to achieve the necessary control and specificity.  In eukaryotic cells, DNA is packaged into highly organized structures called chromosomes. The efficient packaging of long DNA molecules within the limited space of the nucleus necessitates intricate folding and coiling mechanisms. The formation of nucleosomes, chromatin fibers, and higher-order chromosome structures involves precise folding patterns and interactions. The complex packaging of DNA molecules to maintain their stability and accessibility requires sophisticated design and engineering.  The double helical structure of DNA provides stability and protection to the genetic material. It shields the nitrogenous bases within the helix, reducing their exposure to potentially harmful agents that could induce mutations or damage the DNA molecule. The structural integrity of DNA, including the coiling and hydrogen bonding, is finely tuned to resist physical and chemical stresses. The robustness of DNA's design suggests the involvement of an intelligent agent in providing mechanisms for its protection and longevity. When considering the complexity, specificity, precision, and functional integration of the features associated with the coiling of DNA into a double helix, it becomes highly improbable that these characteristics arose solely through unguided processes. The intricate design and interdependent functionalities strongly suggest the involvement of an intelligent agent capable of purposeful design, consistent with what we observe in human-designed systems.

The Role of Cations and Anions in Stabilizing Nucleic Acid Structures

The electrostatic interactions of charged phosphate groups are crucial for the stability of nucleic acid structures. Nucleic acids, such as DNA and RNA, consist of chains of nucleotides, and the negatively charged phosphate groups along the backbone of these chains can repel each other due to their like charges. To counteract this repulsion and maintain the structural integrity of nucleic acids, various cations can interact with the phosphate groups. Cations are positively charged ions that are formed when an atom or a molecule loses one or more electrons. When an atom loses electrons, it becomes positively charged because the number of protons in its nucleus is greater than the number of electrons surrounding it. This positive charge is balanced by the negatively charged electrons that remain in the atom or by other surrounding negatively charged ions. Cations can be formed from various elements across the periodic table. For example, sodium (Na) can lose one electron to form a sodium cation (Na^+), while calcium (Ca) can lose two electrons to form a calcium cation (Ca^2+). The charge on a cation indicates the number of electrons lost by the atom. Monovalent cations, such as Na^+, Li^+, and K^+, are commonly found in biological systems and can electrostatically shield the anionic phosphate groups from each other. These cations interact nonspecifically with the phosphate groups, meaning they do not have a strong preference for binding to a particular location or sequence within the nucleic acid molecule. By surrounding the negatively charged phosphates, monovalent cations effectively reduce the repulsion between adjacent phosphate groups and stabilize the overall structure of the nucleic acid. Monovalent cations, such as Na^+ and K^+, stabilize the overall structure of nucleic acids by reducing the repulsion between adjacent phosphate groups through a process called electrostatic shielding. This shielding effect is a result of the attractive forces between the positively charged cations and the negatively charged phosphate groups. When a monovalent cation approaches a negatively charged phosphate group in a nucleic acid molecule, the positive charge of the cation attracts the negatively charged oxygen atoms in the phosphate group. This attraction partially neutralizes the negative charge on the phosphate group and reduces the electrostatic repulsion between adjacent negatively charged phosphate groups. By surrounding the negatively charged phosphates, monovalent cations form an ionic cloud around the nucleic acid molecule. This cloud of positive charges helps to shield the repulsive forces between adjacent phosphate groups, allowing them to come closer together and maintain a more compact and stable structure. The shielding effect of monovalent cations is nonspecific, meaning they interact with phosphate groups regardless of their specific sequence or location in the nucleic acid molecule. They act as general electrostatic compensators, reducing the overall electrostatic repulsion throughout the nucleic acid structure. While monovalent cations can reduce the repulsion between adjacent phosphate groups and contribute to the stability of nucleic acid structures, they are not as effective as divalent cations like Mg^2+ in this regard. Divalent cations have stronger and more specific interactions with the phosphate groups, leading to even greater stabilization effects.
On the other hand, divalent cations, particularly Mg^2+, Mn^2+, and Co^2+, exhibit specific binding to the phosphate groups in nucleic acids. These cations have a stronger affinity for negatively charged phosphates compared to monovalent cations. Divalent cations form coordination bonds with the phosphate groups, meaning they have a specific arrangement of interactions with the phosphate oxygens. This specific binding provides even stronger shielding and stabilization effects on the nucleic acid structure. Among the divalent cations, magnesium ions (Mg^2+) play a particularly important role in nucleic acid stability. An individual Mg^2+ ion can have an impact on the DNA double helix comparable to that of hundreds to thousands of sodium ions (Na^+). This is because magnesium ions have a higher charge and a more specific interaction with the phosphate groups, allowing them to more effectively shield and stabilize the nucleic acid structure. The significance of magnesium ions extends beyond stability alone. Many enzymes that interact with nucleic acids or catalyze reactions involving nucleotides require magnesium ions as cofactors for their activity. 

Magnesium ions, essential for proper DNA and RNA functioning

The presence of Mg^2+ ions can facilitate the proper folding of RNA molecules, stabilize RNA secondary structures, and participate in RNA-protein interactions. Therefore, magnesium ions are not only important for the stability of nucleic acids but also for the proper functioning of numerous biological processes involving nucleic acids. Cells possess specific transport proteins or ion channels that facilitate the movement of magnesium ions across cell membranes. These transporters allow the entry of extracellular magnesium ions into the cytoplasm or the release of intracellular magnesium ions to the extracellular space. Transporters involved in magnesium ion movement include magnesium transporters, such as TRPM6 and TRPM7, as well as ion channels like MagT1 and SLC41A1. Cells can store magnesium ions within various cellular compartments, such as the endoplasmic reticulum, mitochondria, and vacuoles. These compartments contain magnesium-binding proteins and complexes that sequester and release magnesium ions as needed. Intracellular magnesium levels are tightly regulated to maintain cellular homeostasis. Cells employ various mechanisms to control magnesium ion concentrations. For instance, magnesium ion influx and efflux are regulated by ion transporters, and intracellular magnesium levels can be modulated by magnesium-binding proteins, enzymes, and signaling pathways.  Magnesium ions can also undergo exchange with other ions within cells. For example, magnesium ions can be exchanged with other divalent cations, such as calcium (Ca^2+), through ion transporters. This exchange helps maintain the appropriate balance of magnesium ions and other divalent cations in the cellular environment.

Magnesium Transporters

The presence and proper functioning of these magnesium transporters and ion channels are vital for maintaining the essential levels of magnesium ions required for cellular processes, making them life essential. Magnesium ions play crucial roles in various cellular processes and are involved in numerous biochemical reactions. They regulate enzyme activity, support DNA and RNA synthesis by acting as a cofactor for various enzymes involved in these processes, facilitate muscle contraction, and participate in neurotransmission, among many other functions. The transport proteins and ion channels mentioned, such as TRPM6, TRPM7, MagT1, and SLC41A1, are responsible for the movement of magnesium ions across cell membranes. These transporters ensure that an appropriate balance of magnesium ions is maintained inside and outside the cell, allowing for the proper functioning of cellular processes. Disruptions in magnesium ion homeostasis can have severe consequences on cellular function and overall health. While cells can potentially survive with fluctuations in magnesium levels for a certain period, sustained disruptions in magnesium ion homeostasis can eventually compromise cellular function and viability. 

G. A. C. Franken (2022 Jul 12)   Magnesium (Mg2+) is the most prevalent divalent intracellular cation. As a co-factor in many enzymatic reactions, Mg2+ is essential for protein synthesis, energy production, and DNA stability. Disturbances in intracellular Mg2+ concentrations, therefore, unequivocally result in delayed cell growth and metabolic defects. To maintain physiological Mg2+ levels, all organisms rely on balanced Mg2+ influx and efflux via Mg2+ channels and transporters.

TRPM6 for example are transporters responsible for facilitating the movement of magnesium ions across cell membranes, ensuring an appropriate balance of magnesium ions inside and outside the cell. Magnesium ions act as cofactors for various enzymes involved in DNA and RNA synthesis, protein synthesis, energy production, and other cellular processes. These processes are fundamental to cell viability and function. TRPM6 transporters contribute to maintaining the required levels of magnesium ions, ensuring the availability of this essential cofactor for enzymatic reactions. Disturbances in intracellular magnesium concentrations can lead to delayed cell growth and metabolic defects. TRPM6 transporters help maintain physiological levels of magnesium ions, which are essential for cell growth and division. By facilitating magnesium ion influx, TRPM6 transporters support proper cellular growth and viability.
TRPM6 acts as a channel protein, allowing the passage of magnesium ions across the cell membrane. It is composed of multiple subunits and forms a tetrameric structure, meaning it consists of four individual protein subunits. Each subunit has transmembrane segments that span the cell membrane multiple times, forming a pore or channel through which ions can pass.  TRPM6 channels can be activated by factors such as intracellular magnesium levels, voltage changes across the cell membrane, and phosphorylation events. 

In free-living cells, such as bacteria, the movement of magnesium ions across the cell membrane is also driven by concentration gradients and electrochemical forces. These cells must acquire magnesium from their external environment to fulfill their physiological needs. Bacteria employ various mechanisms to transport magnesium ions into the cell. One common method is through the use of specific transport proteins or channels that facilitate the movement of magnesium ions across the cell membrane. These transport proteins can be categorized into two main types: primary active transporters and secondary active transporters. Primary active transporters utilize energy from ATP hydrolysis to actively transport magnesium ions against their concentration gradient, from areas of lower concentration to areas of higher concentration within the cell. These transporters often have ATP-binding domains and undergo conformational changes to transport magnesium ions across the membrane. Secondary active transporters, on the other hand, harness the energy stored in ion gradients (such as the proton gradient) established by other transport processes in the cell. These transporters use the energy from the established gradients to facilitate the co-transport of magnesium ions along with other ions or molecules across the cell membrane. Additionally, some bacteria may have magnesium channels that allow passive diffusion of magnesium ions down their concentration gradient. These channels are selective for magnesium ions and enable their movement across the membrane without the need for energy input.  Different bacteria may possess different sets of transport proteins or channels to acquire magnesium ions from their surroundings. These mechanisms are adapted to the specific environmental conditions in which the bacteria live, ensuring an adequate supply of magnesium for cellular functions.

There are several different transport proteins that have been identified in bacteria and other organisms for the uptake and transport of magnesium ions. These proteins are often categorized into families or classes based on their structural and functional similarities. Following, a few examples of magnesium transport proteins:

CorA Family: CorA proteins are widespread in bacteria and play a role in magnesium uptake. They are integral membrane proteins that form ion channels and are involved in passive transport of magnesium ions.
MgtE Family: MgtE proteins are found in bacteria and some archaea. They act as magnesium transporters and are important for maintaining intracellular magnesium homeostasis.
MgtA and MgtB: These transporters are present in certain bacteria and are involved in magnesium uptake. They are ATP-driven pumps that actively transport magnesium ions into the cell.
MgtC: MgtC is a membrane protein found in some bacteria, and its exact mechanism is not fully understood. It has been suggested to be involved in magnesium transport or regulation.
MgtR: MgtR is a regulatory protein that controls the expression of magnesium transporters in response to magnesium levels. It is found in certain bacteria and helps regulate magnesium uptake and homeostasis.

These families exhibit structural and functional differences. While they all participate in magnesium transport or regulation, their specific mechanisms and structures can vary. The CorA family proteins are integral membrane proteins that form ion channels. They are involved in passive transport, allowing magnesium ions to move across the cell membrane down their concentration gradient. MgtE proteins, on the other hand, are also involved in magnesium transport but have a different structural organization compared to the CorA family. MgtE proteins are believed to be magnesium-specific transporters and are crucial for maintaining intracellular magnesium homeostasis.
MgtA and MgtB are ATP-driven pumps responsible for active transport of magnesium ions into the cell. These transporters utilize ATP hydrolysis to drive the movement of magnesium against its concentration gradient.
MgtC is a membrane protein with an unclear mechanism. While its exact role in magnesium transport or regulation is not fully understood, it has been suggested to be involved in magnesium transport or regulation.
MgtR is a regulatory protein that controls the expression of magnesium transporters in response to magnesium levels. It acts as a transcriptional regulator, modulating the expression of magnesium transport genes in bacteria.

The presence of different families or types of transporters (CorA, MgtE, MgtA/B, MgtC) that serve the same purpose of magnesium uptake and regulation in different bacteria is an example of convergence. While these proteins may have different structures and mechanisms, they all fulfill the essential function of facilitating magnesium transport. Convergence suggests that there are functional constraints and specific requirements for magnesium transport in different organisms living in diverse environments. The emergence of similar solutions through independent processes points to the existence of design principles that guide the development of effective magnesium transport mechanisms. This implies that the need for magnesium acquisition and homeostasis is a fundamental aspect of life that necessitates purposeful design. Moreover, the convergence of magnesium transport proteins highlights the idea that certain functional requirements, such as maintaining intracellular magnesium levels, are critical for the proper functioning of cells and organisms. The presence of multiple, distinct transport systems that converge on the same function emphasizes the importance and indispensability of magnesium in cellular processes. This convergence strengthens the argument for the existence of a designer who incorporated common design elements across different organisms to achieve similar functional outcomes.

Anions


Anions, such as chloride (Cl-) and other negatively charged ions, also play important roles in stabilizing nucleic acid structures. While cations, particularly magnesium ions (Mg2+), are primarily responsible for neutralizing the repulsion between negatively charged phosphate groups, anions contribute to the overall electrostatic balance and stability of nucleic acids. In the presence of divalent cations like Mg2+, anions help in maintaining the overall charge neutrality of the system. As divalent cations coordinate with the negatively charged phosphate groups, anions provide counterbalancing negative charges to maintain overall charge balance. This balance is crucial for the stability and proper folding of nucleic acid structures. Moreover, anions can also participate in specific interactions with nucleic acids. For example, chloride ions (Cl-) can form ionic interactions with positively charged functional groups, such as amino groups in nucleic acid bases or positively charged residues in proteins. These interactions can contribute to the stability of nucleic acid-protein complexes or facilitate specific protein-nucleic acid interactions.

The presence of cations and anions in nucleic acid structures demonstrates a delicate balance that is necessary for their stability. The specific affinity of divalent cations like magnesium ions (Mg2+) for the negatively charged phosphate groups highlights the fine-tuning required for optimal stabilization. This level of precision suggests intentional design rather than a random implementation. Divalent cations, such as Mg2+, exhibit specific binding to phosphate groups and form coordination bonds, indicating a highly specialized interaction. This specificity allows for stronger shielding and stabilization effects on the nucleic acid structure. The specific interactions between anions and positively charged functional groups further support the idea of purposeful design, as they contribute to the overall stability and specific functions of nucleic acids. The presence and regulation of cations and anions are essential for proper DNA and RNA functioning. Magnesium ions, in particular, are not only crucial for nucleic acid stability but also for various biochemical reactions, enzyme activity, and DNA and RNA synthesis. The intricate system of transporters and channels involved in maintaining appropriate magnesium ion concentrations highlights the importance of these ions in cellular processes. This functional necessity suggests that the presence and regulation of cations and anions are designed to support the complex machinery of life.  The tight regulation of intracellular magnesium ion levels and the involvement of transport proteins, ion channels, and signaling pathways in maintaining cellular homeostasis imply a sophisticated control system. Such a system, which ensures the balance of magnesium ions and other divalent cations, indicates an intelligent design that enables cells to function optimally.

DNAs phosphate ion

To ensure the viability of life's long-term genetic information storage, it is crucial that DNA does not break down in the presence of water. This hydrolysis problem had to be solved beforehand, or else the genetic information would rapidly dissolve, much like a sand castle washed away by the incoming tide. The way DNA addresses this challenge is a remarkable feat of engineering. DNA is a polymeric ester, consisting of a long phosphate (PO₄³-) backbone that stretches close to two meters in humans. This molecular structure is perfectly suited for DNA's purpose. The chemical structure of the phosphate anion, with its four terminal oxygen atoms and three net charges, allows it to bind to two ribonucleotides using two of these oxygen atoms, while one oxygen atom remains single-charged. Represented as (R₁O)(R₂O)P(=O)-O-, where "R" represents a ribonucleotide, this configuration retains a negative charge at the end, which is in resonance with two oxygen atoms. This charge resonance is crucial as it stabilizes the DNA molecule against hydrolysis by water. It forms an electrical shield around the entire DNA double helix, protecting it from reacting with water molecules. Additionally, this encompassing electrical field plays a role in keeping DNA inside the cell nucleus, preventing it from escaping through the cell membrane. These properties make the phosphate anion (PO₄³-) the ideal building block for constructing a stable DNA macromolecule. It bonds well with the appropriate sugars and bases, providing protection against hydrolysis and ensuring the DNA remains encapsulated within the nuclear membrane. To further enable DNA to function properly, another challenge had to be overcome. While inorganic phosphate (PO₄³-) is a suitable link for DNA, its reaction with deoxyribose (a sugar molecule) is naturally slow. Therefore, the cell required a catalyst to accelerate this crucial reaction. Enzymes, large biomolecules with intricate designs, fulfill this role by significantly speeding up the formation of phosphate-sugar bonds by many orders of magnitude. The production of enzymes is a remarkable process in itself, which we will explore later. From the very beginning, enzymes were necessary to create DNA. Yet, they rely on the DNA sequence to produce them. Thus, we have two ingenious solutions to critical challenges: an electrical shield that protects DNA from breaking down in the presence of water, and enzymes that accelerate the formation of phosphate-sugar bonds, a reaction that would otherwise be too slow. These solutions had to be present simultaneously because the DNA sequence is needed to produce the enzymes, while the enzymes are essential for creating DNA. If only one of them existed without the other, no cellular life would be possible.

Topoisomerases

The operation of topoisomerases is truly a mind-boggling and awe-inspiring phenomenon in the biological world. These remarkable molecular machines perform their tasks with extraordinary precision and brilliance, ensuring the flawless untangling of knots within our DNA. DNA replication encounters a critical challenge that must be overcome before it can successfully complete its mission. The separation of the DNA strands leads to twisting in the portion that has not yet been separated. As tension from the twisting increases, the uncopied segment of DNA wraps around itself, forming what we refer to as supercoils. If left unaddressed, these supercoils would impede the DNA replication process, rendering the two strands inseparable and ultimately causing the death of cells. This is where the extraordinary topoisomerases step in to save the day. Topoisomerases are a class of special proteins that possess the ability to untangle knots within DNA. There are two main types of topoisomerases, with type 2 being the more prominent. Type 2 topoisomerases typically consist of three distinct sections: an upper gate, a middle gate, and a lower gate. Each gate can open or close during the operation of the protein. Their exceptional and unparalleled abilities make them a stunning masterpiece of biological ingenuity. Topoisomerases are a group of enzymes that play a crucial role in altering the supercoiling of DNA, maintaining its proper topological state, and facilitating essential biological processes such as replication and transcription. Topoisomerases are essential for life because they play a crucial role in DNA replication. During replication, the DNA double helix needs to unwind and separate into two individual strands to serve as templates for the synthesis of new DNA strands. Topoisomerases help alleviate the tension that builds up ahead of the replication fork by relaxing the supercoils formed during unwinding. This ensures smooth and accurate DNA replication, allowing for the faithful transmission of genetic information to daughter cells. Transcription is the process by which RNA molecules are synthesized using DNA as a template. As RNA polymerase moves along the DNA strand, it generates positive supercoiling ahead of itself. Topoisomerases act to remove these supercoils, preventing the DNA from becoming overly twisted and allowing efficient and continuous transcription. During cell division, chromosomes must be properly segregated into daughter cells. Topoisomerases, particularly type II topoisomerases, play a vital role in chromosome segregation. They help resolve the intertwining of DNA strands and decatenate replicated chromosomes by introducing transient double-strand breaks. This ensures that each daughter cell receives an accurate and complete set of chromosomes.  Topoisomerases are involved in DNA repair processes. They assist in the repair of DNA damage by manipulating the topological state of DNA. For example, when DNA strands become tangled or knotted due to DNA damage, topoisomerases can introduce and resolve DNA strand breaks to untangle the DNA and restore its proper structure. Topoisomerases contribute to the regulation of gene expression. By altering the supercoiling of DNA, they influence the accessibility of specific regions of DNA to transcription factors and other regulatory proteins. Changes in DNA supercoiling can modulate gene expression by facilitating or hindering the binding of regulatory proteins to DNA, thus controlling the activation or repression of genes.

The process by which type 2 topoisomerases untangle DNA can be summarized in four main steps:

1. Two DNA segments enter through the top gate, and the middle gate is used to break one segment of DNA apart.
2. The second DNA segment is then passed through the break, crucially untangling the two segments.
3. The topoisomerase recombines the first DNA segment, resulting in the elimination of two supercoils.
4. Finally, the untangled DNA segments are released, with the second strand released at the bottom and the first strand released at the top.

This summary represents a simplified version of the process, and the actual mechanism is much more complex. Recent research has shed light on the intricate details of how this process occurs. When we zoom in on the upper part of the topoisomerase, we observe that one of the two overlapping DNA segments enters the upper gate and binds to the middle gate. Subsequently, the second DNA segment also enters the upper gate, and the topoisomerase cuts the DNA in the middle gate into two pieces. Two ATP molecules attach to the upper gate, causing it to close. The breaking apart of one ATP molecule releases energy in the form of adenosine diphosphate (ADP) and phosphate, which helps maintain the closure of the upper gate during subsequent steps. The middle gate then opens, pulling the broken halves of DNA apart and creating a gap. As the middle gate remains connected to the broken ends, the DNA remains attached to it. The second DNA segment moves through this gap, after which the middle gate closes, reconnecting the broken ends of the DNA. The upper gate rotates, preventing the second segment from reversing through the break. Finally, the remaining ATP molecule breaks apart, leading to the opening of the lower gate, allowing the second DNA segment to leave. Subsequently, the lower gate closes, the upper gate opens, and the first DNA segment is released. Once this process is complete, the topoisomerase is reset and ready to repeat the same sequence of steps. The topoisomerase molecular machine exemplifies the wonders of the biological world, exhibiting an intricate and intelligent orchestration of its operation. The operation of topoisomerases is a testament to the remarkable design of these molecular machines. Their ability to open and close gates, cut and rejoin DNA strands, and utilize ATP molecules to generate energy showcases the sophistication of their mechanisms. These processes occur with remarkable precision, ensuring the successful resolution of DNA supercoils and the continuation of DNA replication. Moreover, the role of topoisomerases extends beyond DNA replication. They play crucial roles in various cellular processes, such as transcription, where DNA is used as a template to produce RNA molecules. Topoisomerases help unwind DNA strands to facilitate the transcription process, allowing the genetic information to be transcribed accurately. Additionally, topoisomerases are involved in DNA repair mechanisms. When DNA strands are damaged, they can form knots and tangles that hinder proper repair processes. Topoisomerases act as guardians, untangling these knots and enabling efficient DNA repair, thereby preserving the integrity of our genetic material. The intricate and intelligent operation of topoisomerases highlights the extraordinary nature of biological systems and their ability to solve complex challenges. Scientists continue to explore and unravel the detailed mechanisms and regulation of topoisomerases, enhancing our understanding of these remarkable molecular machines. Inspired by the ingenuity of topoisomerases, researchers are also investigating ways to harness their capabilities for practical applications. Understanding and manipulating these processes could lead to the development of novel therapeutic approaches for various diseases, including cancer. Targeting topoisomerases could potentially disrupt the replication and repair processes in cancer cells, providing new avenues for treatment.

There are two classes of topoisomerases: Type I and Type II, found in both prokaryotes and eukaryotes.

Type I topoisomerases create transient single-strand breaks in DNA to alter its supercoiling. Type I enzymes are further classified into two subtypes: type IA and type IB. Type IA topoisomerase enzymes are present in all cells and specifically relax negatively supercoiled DNA. They operate by cutting a single strand of DNA, passing a single-strand loop through the resulting gap, and then resealing the break. This process increases the linking number and reduces the supercoiling of the DNA molecule. Type IB topoisomerase enzymes also relax negatively supercoiled DNA but have a different sequence and reaction mechanism compared to type IA topoisomerases. They perform similar strand passage mechanisms to alter the supercoiling of DNA. Type II topoisomerase enzymes create transient double-strand breaks in DNA to alter its supercoiling. Unlike Type I topoisomerases, they act on both negatively and positively supercoiled DNA. Type II topoisomerases are involved in processes such as DNA replication, chromosome segregation, and recombination. They use ATP hydrolysis to pass one DNA segment through another, thereby altering the supercoiling and topology of the DNA molecule. The relaxation of negatively supercoiled DNA by type IA topoisomerases occurs through a strand passage mechanism. These enzymes cut a single DNA strand, pass a single-strand loop through the gap, and then reseal the break. This process increases the linking number and reduces the supercoiling of the DNA molecule.

The structure of type IIA topoisomerases consists of several key motifs that are crucial for their function:

N-terminal GHKL ATPase Domain: This domain is responsible for ATP hydrolysis and provides the energy required for the enzyme's catalytic activity. The GHKL domain forms a dimeric structure that encloses ATP within its active site.

Toprim Domain: The Toprim domain is a Rossmann fold that contains three invariant acidic residues coordinating magnesium ions involved in DNA cleavage and religation. It is involved in DNA binding and recognition, as well as catalytic activity. The catalytic tyrosine, which plays a central role in the cleavage and rejoining of DNA strands, is located within this domain.

Central DNA-binding Core: The central core of the enzyme contains the Toprim fold and a DNA-binding core. The DNA-binding core consists of a winged helix domain (WHD) or CAP domain, which resembles the WHD of catabolite activator protein. The WHD leads to a tower domain, followed by a coiled-coil region that links to the C-terminal domain.

C-terminal Domain: The C-terminal domain forms the main dimer interface for the enzyme. It is involved in stabilizing the overall structure of the enzyme and maintaining the integrity of the catalytic site.

The ATPase domains of the enzyme dimerize to form a closed conformation, enclosing ATP within the active site. The transducer domain, linking the ATPase domain to the Toprim fold, is believed to transmit the nucleotide state of the ATPase domain to the rest of the protein, coordinating the catalytic cycle. The structure of the enzyme reveals that the DNA is bent by approximately 160 degrees through an invariant isoleucine residue. This bending is critical for the proper positioning and interaction of the enzyme with DNA during the catalytic process.  The specific amino acids involved in catalysis are responsible for several critical functions. Amino acids within the DNA-binding core, such as those within the winged helix domain (WHD), interact with the DNA molecule, ensuring proper recognition and binding of the substrate. These interactions involve both electrostatic interactions, such as hydrogen bonding and ionic interactions, as well as van der Waals forces. The winged helix domain (WHD), also known as a forkhead domain or winged helix-turn-helix domain, is a common structural motif found in many DNA-binding proteins, including type IIA topoisomerases. It plays a crucial role in ensuring proper recognition and binding of the DNA substrate. The WHD consists of three α-helices (H1, H2, and H3) and a three-stranded β-sheet, with two short helical segments called wings (W1 and W2) extending from the β-sheet. The WHD forms a compact fold, with the wings positioned above the DNA-binding surface. The DNA-binding surface of the WHD contains a series of positively charged residues, such as arginine and lysine, which can form favorable electrostatic interactions with the negatively charged phosphate backbone of DNA. This electrostatic interaction helps stabilize the protein-DNA complex. Cleavage and rejoining of DNA strands: The catalytic tyrosine residue within the Toprim domain plays a crucial role in the cleavage and rejoining of DNA strands. This tyrosine residue undergoes a transient covalent attachment to the DNA backbone during the reaction, facilitating the strand breakage and rejoining processes. Amino acids within the ATPase domain are involved in coordinating ATP and facilitating its hydrolysis, providing the energy required for the enzyme's catalytic cycle. These amino acids contribute to the proper positioning and orientation of ATP within the active site. The precise rotation angle of atoms within certain amino acids can indeed be crucial for the catalytic activity of enzymes. The three-dimensional arrangement of atoms within the active site determines the specific interactions between the enzyme and its substrate. Small changes in the rotation angles of atoms can significantly affect the ability of the enzyme to bind to its substrate, form the necessary catalytic intermediates, and facilitate the reaction.



The catalytic cycle of topoisomerase II involves several stages:

DNA Binding: The enzyme binds to the DNA molecule, recognizing and interacting with specific DNA sequences. The ATPase domain interacts with the DNA to facilitate proper positioning and stabilization of the enzyme on the DNA substrate.
ATP Hydrolysis: The ATPase domain utilizes ATP hydrolysis to provide the energy required for the enzyme's catalytic activities. ATP binding induces conformational changes in the enzyme, leading to the formation of a closed clamp around the DNA.
DNA Cleavage: One DNA strand, referred to as the gate (G) strand, is cleaved by the enzyme. This cleavage creates a transient DSB in the DNA helix. The cleavage domain (B) of topoisomerase II is responsible for making the DNA strand break.
DNA Strand Passage: Another intact DNA strand, referred to as the transport (T) strand, passes through the DNA break site. This passage allows for the alteration of DNA supercoiling and topology.
DNA Religation: After the strand passage, the DNA strand break is resealed, restoring the continuity of the DNA helix. The cleavage domain (B) catalyzes the religation of the DNA strands.

The catalytic cycle of topoisomerase II is a highly specialized and intricate process that enables the enzyme to perform its essential role in DNA metabolism. Each step of the catalytic cycle, from DNA binding to ATP hydrolysis, DNA cleavage, DNA strand passage, and DNA religation, is interconnected and dependent on the successful completion of the previous steps. If any of these steps were missing or implemented out of order, the enzyme would not be able to carry out its function effectively. In other words, a designer had to design and implement all the necessary components and mechanisms of the catalytic cycle simultaneously for the enzyme to be functional right from the start. A stepwise implementation would not be productive because intermediate stages would lack the necessary coordination and integration required for the enzyme to perform its intended function. Therefore, the catalytic cycle of topoisomerase II had to be implemented as a complete and functional system from its inception, ensuring its ability to fulfill its role in DNA metabolism. The recognition and binding of DNA by the WHD involve several key steps:  The WHD has a specific binding pocket formed by amino acid residues within the domain. These residues can interact with specific DNA sequences or motifs through hydrogen bonding and van der Waals interactions. This sequence-specific recognition allows the protein to selectively bind to its target sites on DNA.  The overall shape of the WHD and its wings allow it to fit into the major groove of the DNA helix. The wings contribute to the binding specificity by making additional contacts with the DNA bases in the major groove. This shape complementarity enhances the stability of the protein-DNA complex and ensures proper binding.  Upon binding to DNA, the WHD can undergo conformational changes to optimize its interaction with the DNA substrate. This induced fit mechanism helps to further stabilize the protein-DNA complex and improve binding affinity. The binding of the WHD to DNA can induce local distortions in the DNA structure. These distortions can involve bending or unwinding of the DNA helix, which can facilitate other steps in the catalytic cycle of the topoisomerase enzyme.

The emergence of the specific steps involved in the recognition and binding of DNA by the WHD (winged-helix domain) of topoisomerase II on a prebiotic Earth through random processes is highly unlikely. The WHD of topoisomerase II possesses a specific binding pocket that interacts with DNA sequences or motifs through precise hydrogen bonding and van der Waals interactions. The chance of such a specific pocket evolving randomly on a prebiotic Earth is extremely low. Specificity requires a precise arrangement of amino acid residues within the domain, which is improbable to occur unless it was designed.  The overall shape of the WHD and its wings allows it to fit into the major groove of the DNA helix. This shape complementarity enhances the stability of the protein-DNA complex and ensures proper binding. Achieving such complementary shapes between the protein and DNA purely by random chance would be highly unlikely. Upon binding to DNA, the WHD can undergo conformational changes to optimize its interaction with the DNA substrate. This induced fit mechanism helps to further stabilize the protein-DNA complex and improve binding affinity. The ability of the WHD to undergo specific conformational changes in response to DNA binding is a sophisticated feature that is unlikely to arise randomly. The binding of the WHD to DNA can induce local distortions in the DNA structure, such as bending or unwinding of the DNA helix. These distortions facilitate other steps in the catalytic cycle of topoisomerase II. The emergence of such specific distortions in DNA structure through random processes is highly improbable. The topoisomerase II enzyme would have no function without a fully operating genome and DNA replication. This enzyme plays a critical role in DNA metabolism, particularly in managing DNA supercoiling and topology. It is part of a complex network of cellular processes that involve DNA replication, transcription, and repair. The presence of a fully functional genome and the associated cellular machinery is necessary for the topoisomerase II enzyme to have any meaningful function and impact on cellular processes. The conceptual paradox of DNA replication and the need for topoisomerase adds another layer of complexity. DNA needs topoisomerase to unwind itself for replication, but in order to pass on the genes coding for topoisomerase, it would already require topoisomerase. This scenario implies that DNA must code for the protein itself, which raises questions about how this code could have emerged in the first place. The mechanics of unwinding, cutting, and re-joining DNA sections require precise biochemistry, further adding to the challenges.  The wider issue of DNA replication involves a multitude of interconnected processes and proteins. Proteins that act as stabilizing clamps, DNA unzipping enzymes, error-checking mechanisms, and the intricate duplication of one DNA strand backwards and in sections are just a few examples of the complexity involved. Explaining the origin and coordination of all these components through gradual, step-by-step evolutionary processes presents significant hurdles.



The X-ray structures of yeast topoisomerase II provide valuable insights into the organization and functioning of this enzyme. The structures reveal two key domains involved in the catalytic cycle: the N-terminal ATPase domain and the DNA breakage/reunion domain.

(a) N-terminal ATPase domain: This domain, comprising residues 7-406 of the subunits, forms a homodimer. In the X-ray structure, the protein is represented as a ribbon, with one subunit shown in gray and the other in rainbow order, indicating a color gradient from N-terminus (blue) to C-terminus (red). The homodimer is arranged with a twofold axis, implying that the two subunits have a similar structure but with an unknown relative orientation about the axis. The X-ray structure also reveals the presence of AMPPNP molecules, depicted in space-filling form, bound to the ATPase domain. The AMPPNP molecules are shown in green (carbon atoms), blue (nitrogen atoms), red (oxygen atoms), and orange (phosphorus atoms). These molecules represent the non-hydrolyzable ATP analog and provide insights into the ATPase activity of the enzyme.

(b) DNA breakage/reunion domain: This domain, comprising residues 419-1177, also forms a homodimer. The X-ray structure shows the domain in complex with a doubly nicked 34-bp DNA molecule. The DNA molecule is represented as a semitransparent molecular surface in orange, with the atoms colored according to the standard convention (carbon in green, nitrogen in blue, oxygen in red, and phosphorus in orange). The structure of the DNA molecule describes a 150° arc. The protein subunits are depicted as ribbons, with one subunit in gray and the other in rainbow order, indicating the color gradient from N-terminus (blue) to C-terminus (red). Notably, the X-ray structure highlights the active site residues, particularly the two active site Tyr residues (Y782), which are shown in space-filling form in magenta (carbon atoms) and red (oxygen atoms). These residues play a critical role in catalyzing the cleavage and rejoining of DNA strands.

The X-ray structures provide a detailed view of the interactions between the protein domains and the DNA substrate, shedding light on the mechanisms underlying the catalytic activities of topoisomerase II. The structures also offer valuable information about the overall organization and spatial arrangement of the protein domains, allowing for a better understanding of how they cooperate during the catalytic cycle.

Nucleic acids rely on base pairing, stacking, and ionic interactions for stability

Nucleic acids, such as DNA and RNA, are stabilized by various forces, including base pairing, stacking interactions, and ionic interactions. These forces contribute to the overall stability and structural integrity of nucleic acid molecules.
Base pairing is a fundamental mechanism that holds together the complementary strands of double-stranded nucleic acids. In DNA, the Watson-Crick base pairs are the most common and stable type of base pairing. Adenine (A) forms hydrogen bonds with thymine (T) using N1 and N3 as hydrogen bond acceptors, while guanine (G) forms hydrogen bonds with cytosine (C) using N1 and O2 as acceptors. The specific hydrogen bonding patterns between the bases create a tight and specific pairing that contributes to the stability of the double helix.  While non-Watson-Crick base pairs are theoretically possible, experimental observations have shown that Watson-Crick geometry is the most stable mode of base pairing in the double helix. This stability arises from the intrinsic affinity between the bases in a Watson-Crick pair, which is higher compared to non-Watson-Crick pairs. The geometric constraints of the double helix also contribute to the prevalence of Watson-Crick base pairs. The similar shapes of A-T and G-C base pairs allow for their interchange without altering the conformation of the sugar-phosphate backbone, maintaining the overall structure of the double helix.

Nucleic acids are carriers of genetic information and have a high degree of specificity and complexity. The precise arrangement of nucleotide bases in DNA and RNA is essential for encoding and transmitting genetic instructions. The probability of randomly assembling the correct sequence of bases necessary for functional genetic information is astronomically low. Base pairing in nucleic acids follows strict rules, such as the Watson-Crick pairing in DNA. Adenine pairs with thymine and guanine pairs with cytosine through specific hydrogen bonding patterns. The precise complementarity and specificity required for stable base pairing make it highly improbable to occur by chance alone. Nucleic acids exhibit homochirality, meaning that their building blocks (sugars and amino acids) exist predominantly in either the right-handed or left-handed form. Achieving homochirality through random processes is extremely unlikely, yet it is crucial for the proper functioning of nucleic acids. Nucleic acids require a specific order of nucleotide incorporation during their assembly. Random assembly would result in a non-functional, random sequence rather than the specific and functional sequences found in DNA and RNA. The synthesis and replication of nucleic acids involve numerous enzymes and molecular machinery, such as DNA polymerases and helicases. These complex systems are required to ensure accurate replication and maintenance of genetic information. The simultaneous emergence of these intricate molecular systems through random processes is highly improbable. The formation and stability of nucleic acids depend on specific environmental conditions, including pH, temperature, and the availability of specific ions. These conditions need to be precisely controlled and maintained for nucleic acids to form and function properly. Achieving such a fine-tuned environment by chance in a prebiotic Earth is highly unlikely.

On one hand, enzymes and molecular machinery are necessary for the accurate replication and maintenance of genetic information. Without them, the replication process would be error-prone and inefficient, leading to the loss or corruption of genetic information over time. These complex systems provide the necessary mechanisms for unwinding the DNA strands, copying the genetic code, proofreading for errors, and repairing any damage. On the other hand, genetic information itself is needed to code for the production of these enzymes and molecular machinery. The instructions for building these complex systems are encoded in the DNA or RNA molecules. Without genetic information specifying the sequences and structures of these molecular components, the enzymes, and machinery required for replication cannot be produced. This presents a dilemma: the accurate replication and maintenance of genetic information require complex systems, but the existence of these systems depends on the genetic information itself. It becomes a circular problem where one component relies on the other for its existence. The intricate and interdependent nature of nucleic acids, their base pairing specificity, the requirement for homochirality, the involvement of complex enzymatic systems, and the circular relationship between genetic information and the production of molecular machinery strongly suggest the involvement of intelligent design as the best explanation for their origin and complexity.

Much has been investigated, written, and said, in the last 70 years, since the advent of Origin of Life research, and about the prebiotic Origin of RNA and DNA. It gave rise to hypotheses, like the RNA world, which is maybe, besided the metabolism first scenario, still today the predominant naturalistic explanation for the Origin of Life. But besides the naturalistic origin of RNA prebiotically, and its non-enzymatic catalysis, polymerization, self-replication, and the question of the Origin of Biological Information, genetic language, and the genetic code, very little has been said about the origin of the metabolic systems, that synthesize RNA and DNA, and their origin, upon which the first living Cell depended upon. In fact, researching a bit of the existing literature, the description of the pathways that lead to the synthesis of these essential building blocks of life is rarely found in popular essays, news reports, or science books, maybe because it gets very technical, and aside of professionals, Chemists, and Biochemists, that work in the field, there is little interest by the general public, to read about this stuff. The description of these biochemical systems is mostly found in biochemistry textbooks, and science papers, read and studied by those working in the field. Usually, in these texts, the authors do little care about the technical jargon and presuppose that the readers have the necessary background knowledge, to understand what is being described and said. Despite having no formal education, I have dived into the literature in the last over ten years, and collected a substantial amount of information, with the goal of elucidating, if the evidence brought to light by science leads to naturalistic explanations, or if intelligent agency as a must, and being required, to explain the origin of these systems in an adequate manner. I have come to a clear conclusion, that is exposed in this book. As Louis Pasteur said:'A bit of science distances one from God, but much science nears one to Him.' I agree with Pasteur, and that is the focus on this book: To elucidate, why, in my opinion, design clearly tops non-design in the explanatory power of these molecular nanosystems. My first book on the Origin of Life: On the Origin of Life and Virus World by Means of an Intelligent Designer: The Factory Maker, Paley's Watchmaker Argument 2.0, dealt mostly with the question of the prebiotic origin of the building blocks of life. This book looks closely into the opposite of the spectrum: The advanced synthesis of RNA and DNA in the first life form, the so-called Progenote, the First Universal Common Ancestor, or, why not Last Universal Common Ancestor. Truth said, nobody knows exactly, what any of those supposed life forms looked like, and what they were made of, to start life. Comparing ten scientific articles related to investigating how LUCA, ( the last universal common ancestor ), there will be ten different answers with a list of life-essential components, systems, and parts. But there is unanimity about the fact, that the LUCAs information system was based on RNA and DNA, and required the metabolic pathways to produce these building blocks. And this is what this book is about. We will delve into these metabolic pathways, the proteins and enzymes, that are essential in these pathways. While, commonly, biochemistry books spend just a few pages to describe these systems, I am dedicating this entire book to looking deeply into these metabolic systems, and each of the enzymes they are composed of, their interconnectedness, their composition, their enzymatic reactions, what co-factors are needed as substrates to feed them, and how these co-factors - in many cases - are synthesized or recruited, the transport proteins, that feed these metabolic pathways with the substrates, the cellular organization, etc. This is a fascinating world, with plenty of scientific literature available, but rarely made available to the public, and written in a simplified language, that can be understood as well by those, not initiated into the technical language. Admittedly, it is a hard task, to write a book, and simplify it enough that the average reader, that has no formal education in the field, can understand and appreciate the description of these systems, but not too simple, to remove the description to be detailed enough, keeping what is so fascinating about this world. Its sophistication, ingeniosity, and awe-inspiring complexity. It was my aim, as far as possible, to explain and describe the technical terms, so that the reader can familiarize themselves with them, and have a learning experience. For me, writing this book, has been an awe-inspiring journey, and i hope, it will be so as well for you, dear reader. But the principal goal is to demonstrate, how the nano-world confirms today, even far more, than 160 years ago, in the times of Louis Pasteur, that science leads to God. In this case, that microbiology leads to God. What is, is the path to the cause. RNA, and DNA, lead to an intelligent designer as the best explanation for the implementation of these super sophisticated molecular production lines.

Over the past 70 years, since the emergence of Origin of Life research, significant progress has been made in understanding the prebiotic origin of RNA and DNA, giving rise to compelling hypotheses such as the RNA world. While much attention has been given to the naturalistic origin of RNA and its vital functions in catalysis, polymerization, and self-replication, relatively little has been said about the origin of the metabolic systems responsible for synthesizing RNA and DNA—the very foundations upon which the first living cell depended.

In popular essays, news reports, and science books, discussions about the pathways leading to the synthesis of these essential building blocks of life are often scarce. This may be attributed to the technical nature of the subject matter, which tends to limit the interest of the general public. Descriptions of these biochemical systems are typically found in specialized biochemistry textbooks and scientific papers, assuming readers possess the necessary background knowledge to comprehend the intricacies involved.

Despite lacking formal education, I have immersed myself in the scientific literature over the past decade, accumulating a wealth of information in an attempt to elucidate whether the evidence presented by science supports naturalistic explanations or necessitates the presence of an intelligent agency to account for the origin of these complex systems. This book represents the culmination of my endeavors and aims to explore why, in my view, design surpasses non-design in its explanatory power concerning these molecular nanosystems.

As Louis Pasteur famously stated, "A bit of science distances one from God, but much science nears one to Him." I wholeheartedly concur with Pasteur, and it is precisely this sentiment that drives the focus of this book: to elucidate why, in my opinion, design unequivocally triumphs over non-design in comprehensively explaining the synthesis of RNA and DNA, and the intricate processes that accompanied the emergence of life.

My previous book, "On the Origin of Life and Virus World by Means of an Intelligent Designer: The Factory Maker, Paley's Watchmaker Argument 2.0," primarily explored the prebiotic origins of life's building blocks. In this current work, we delve into the opposite end of the spectrum, examining the advanced synthesis of RNA and DNA within the first life form—the so-called Progenote, or Last Universal Common Ancestor (LUCA). Although the exact nature and composition of these ancient life forms remain unknown, there is consensus within the scientific community that LUCA's information system relied on RNA and DNA and necessitated metabolic pathways to produce these essential building blocks. It is precisely this aspect that we shall explore in great detail.

While scientific literature presents a multitude of perspectives regarding the components, systems, and parts essential for LUCA, there is unanimity surrounding the significance of RNA and DNA in its information system. This book aims to illuminate the intricate metabolic pathways, the proteins and enzymes vital to these pathways, their interconnectedness, composition, enzymatic reactions, the substrates required as co-factors, and how these co-factors are synthesized or recruited. Additionally, we will investigate the transport proteins responsible for feeding these metabolic pathways with substrates, as well as the underlying cellular organization.

Within this fascinating world, there exists a wealth of scientific literature, often inaccessible to the general public due to its technical language. My goal was to simplify and present this knowledge in a manner that is both understandable and captivating to readers without formal education in the field. While challenging, I endeavored to strike a balance between simplification and retaining the essence and intricacy of these systems—their sophistication, ingenuity, and awe-inspiring complexity. It is my intention to provide an enlightening experience that familiarizes readers with the technical terms and instills a sense of wonder.

Writing this book has been an awe-inspiring journey for me, and I sincerely hope that it proves to be equally captivating for you, dear reader. Above all, the primary objective is to demonstrate how the nano-world, even more so than in the time of Louis Pasteur, unequivocally supports the notion that science points towards a divine creator. In this case, microbiology serves as a pathway to understanding the cause behind the existence of RNA and DNA—the molecular production lines that are undeniably the handiwork of an intelligent designer.