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6.7: Enzymatic Reaction Mechanisms

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  • We can apply what we learned about catalysis by small molecules to enzyme-catalyzed reactions. To understand the mechanism of an enzyme-catalyzed reaction, we try to alter as many variables, one at a time, and ascertain the effects of the changes on the activity of the enzyme. Kinetic methods can be used to obtain data from which inferences about the mechanism can be made. Obviously, crystal structures of the enzyme in the presence and absence of a competitive inhibitor give abundant information about possible mechanisms. It is amazing, however, how much information about enzyme mechanism can be gained even if all you have is a blender, a stopwatch, an impure enzyme, and a few substrates and inhibiting reagents. Systematically, the kineticist, medicinal chemist and molecular biologists (i.e. a well trained chemist) can change:

    1. the substrate - for example, changing the leaving group or acyl sustituents of a hydrolyzable substrate;
    2. the pH or ionic strength - which can give data about general acids/bases in the active site;
    3. the enzyme - by chemical modification of specific amino acids, or through site-specific mutagenesis;
    4. the solvent - as will be discussed in the next chapter section .

    We will explore in detail the mechanisms of three enzymes. For carboxypeptidase, we will study possible mechanisms for the cleavage of C-terminal hydrophobic amino acids from a peptide. For lysozyme, we will study the structural features of the enzyme and substrates along with the mechanism for cleavage of glycosidic links in bacterial peptidoglycan cell walls. For chymotrypsin, we will study experiments which vary the substrate, pH, and the enzyme and infer from this information about a mechanism consistent with the experimental data. Kinetic analyses can be used to determine the:

    • order of binding/dissociation of substrates and products
    • rate constants for individual steps
    • and clues to the nature of catalytic groups found in the enzyme.

    1. Carboxypeptidase

    A peptide substrate binds at the active site of the enzyme. X-ray structures of the enzyme with and without a competitive inhibitor show a large conformational change at the active site when inhibitor or substrate is bound. Without inhibitor, several waters occupy the active site. When inhibitor and presumably substrate are bound, the water leaves (which is entropically favored), and Tyr 248 swings around from near the surface of the protein in the absence of a molecule in the active site to interact with the carboxyl group of the bound molecule, a distance of motion equal to about 1/4 the diameter of the protein. This effectively closes off the active site and expels the water. A Zn2+ ion is present at the active site. It is bound by His 69, His 196, Glu 72, and finally a water molecule as the fourth ligand. A hydrophobic pocket which interacts with the phenolic group of the substrate accounts for the specificity of the protein.

    In the catalytic mechanism, Zn2+ helps polarize the labile amide bond, while Glu 270, acting as a general base, which along with Zn2+ helps promote dissociation of a proton from the bound water, making it a better nucleophile. Water attacks the electrophilic carbon of the sessile bond, with Glu 270 acting as a general base catalyst. The tetrahedral intermediate then collapses, expelling the leaving amine group, which picks up a proton from Glu 270, which now acts as a general acid catalyst. People used to believe that Tyr 248 acted as a general acid, but mutagenesis showed that Tyr 248 can be replaced with Phe 248 without significant effect on the rate of the reaction.

    Jmol: Updated Carboxypeptidase A Jmol14 (Java) | JSMol (HTML5)



    This enzyme, found in cells and secretions of vertebrates but also in viruses which infect bacteria, cleaves peptidoglycan GlcNAc (β 1->4) MurNAc repeat linkages (NAG-NAM) in the cell walls of bacteria and the GlcNAc (β 1->4) GlcNAc (poly-NAG) in chitin, found in the cells walls of certain fungi. Since these polymers are hydrophilic, the active site of the enzyme would be expected to contain a solvent-accessible channel into which the polymer could bind. The crystal structures of lysozyme and complexes of lysozyme and NAG have been solved to high resolution. The inhibitors and substrates form strong H bonds and some hydrophobic interactions with the enzyme cleft. Kinetic studies using (NAG)n polymers show a sharp increase in kcat as n increases from 4 to 5. The kcat for NAG6 and (NAG-NAM)3 are similar. Models studies have shown that for catalysis to occur, (NAG-NAM)3 binds to the active site with each sugar in the chair conformation except the fourth which is distorted to a half chair form, which labilizes the glycosidic link between the 4th and 5th sugars. Additional studies show that if the sugars that fit into the binding site are labeled A-F, then because of the bulky lactyl substituent on the NAM, residues C and E can not be NAM, which suggests that B, D and F must be NAM residues. Cleavage occurs between residues D and E.

    A review of the chemistry of glycosidic bond (an acetal) formation and cleavage shows the acetal cleavage is catalyzed by acids and proceeds by way of an oxonium ion which exists in resonance form as a carbocation.

    Catalysis by the enzyme involves Glu 35 and Asp 52 which are in the active site. Asp 52 is surrounded by polar groups but Glu 35 is in a hydrophobic environment. This should increase the apparent pKa of Glu 35, making it less likely to donate a proton and acquire a negative charge at low pH values, making it a better general acid at higher pH values. The general mechanism appears to involve:

    • binding of a hexasaccharide unit of the peptidoglycan with concomitant distortion of the D NAM.
    • protonation of the sessile acetal O by the general acid Glu 35 (with the elevated pKa), which facilitates cleavage of the glycosidic link and formation of the resonant stabilized oxonium ion.
    • Asp 52 stabilizes the positive oxonium through electrostatic catalysis. The distorted half-chair form of the D NAM stabilizes the oxonium which requires co-planarity of the substituents attached to the sp2 hybridized carbon of the carbocation resonant form (much like we saw with the planar peptide bond).
    • water attacks the stablized carbocation, forming the hemiacetal with release of the extra proton from water to the deprotonated Glu 35 reforming the general acid catalysis.

    Binding and distortion of the D substituent of the substrate (to the half chair form as shown above) occurs before catalysis. Since this distortion helps stabilize the oxonium ion intermediate, it presumably stabilizes the transition state as well. Hence this enzyme appears to bind the transition state more tightly than the free, undistorted substrate, which is yet another method of catalysis.

    pH studies show that side chains with pKa's of 3.5 and 6.3 are required for activity. These presumably correspond to Asp 52 and Glu 35, respectively. If the carboxy groups of lysozyme are chemically modified in the presence of a competitive inhibitor of the enzyme, the only protected carboxy groups are Asp 52 and Glu 35.

    In an alternative mechanism, Asp 52 acts as a nucleophilic catalysis and forms a covalent bond with NAM, expelling a NAG leaving group with Glu 35 acting as a general acid. This alternative mechanism also is consistent with other β-glycosidic bond cleavage enzyme. Substrate distortion is also important in this alternative mechanism.

    Figure: alternative mechanism



    Chymotrypsin, a protease, cleaves amides as well as small ester substrates after aromatic residues. The following data using different chymotrypsin substrates suggests a covalent intermediate occurs on chymotrypsin catalyzed cleavage of esters and amides.

    1. changing the substrate - for example changing the leaving group or acyl sustituents of a hydrolyzable substrate:
    Chymotrypsin substrate cleavage, 25oC, pH 7.9
    kinetic constants Acetyl-Tyr-Gly-amide Acetyl-Tyr-O Ethylester Ester/Amide
    kcat (s-1) 0.50 193 390
    Km (M) 0.023 0.0007 0.03
    kcat/Km (M-1s-1) 22 280,000 12,700

    Kinetic constants for chymotrypsin cleavage of N-acetyl-L-Trp Derivatives - N-acetyl-L-Trp-X

    X kcat (s-1) Km x 103 (M)
    -OCH2CH3 27 0.097
    -OCH3 28 0.095
    -p-nitrophenol 31 0.002
    -NH2 0.026 7.3
    1. the kcat and kcat/Km are larger and the Km smaller for ester substrates compared to amide substrates, suggesting that amides are more difficult to hydrolyze (tables 1 and 2 above). This is expected given the poorer leaving group of the amide.
    2. the kcat for the hydrolysis of ester substrates doesn't depend on the nature of the leaving group (i.e. whether it is a poorer leaving group such as methoxy or a better leaving group such as p-nitrophenolate) suggesting that this step is not the rate limiting step for ester cleavage. (Table 2). Without the enzyme, p-nitrophenyl esters are cleaved much more rapidly than methyl esters. Therefore deacylation must be rate limiting. But deacylation of what? If water was the nucleophile, release of the leaving group would result in both products, the free carboxyl group and the amine being formed simultaneously. This suggests an acyl-enzyme covalent intermediate.
    3. When the acyl end of the ester substrate is changed, without changing the leaving group (a p-nitrophenyl group), a covalent intermediate can be trapped. Specifically, the deacylation of a trimethyacetyl group is much slower than an acetyl group. It is so slow that a 14C-labeled trimethylacetyl-labeled chymotrypsin intermediate can be isolated after incubation of chymotrypsin with 14C-labeled p-nitrophenyltrimethylacetate using gel filtration chromatography.

    We have seen a kinetic mechanism consistent with these ideas before. The reaction equations are shown below:

    In this reaction, a substrate S might interact with E to form a complex, which then is cleaved to products P and Q. Q is released from the enzyme, but P might stay covalently attached, until it is expelled. This conforms exactly to the mechanism described above. For chymotrypsin-catalyzed cleavage, the step characterized by k2 is the acylation step (with release of the leaving group such as p-nitrophenol in Lab 5). The step characterized by k3 is the deacylation step in which water attacks the acyl enzyme to release product P (free phosphate in Lab 5). In class and for homework you derived the following equation::

    Equation 7: v = [(k2k3)/(k2 + k3)]EoS/[Ks(k3)/(k2+k3)] + S

    As mentioned above, for hydrolysis of ester substrates, which have better leaving groups compared to amides, deacylation is rate limiting, ( k3<<k2). Then equation 7 becomes

    v = k3EoS/[Ks(k3)/(k2) + S]

    Vm = k3Eo and Km = Ks(k3)/(k2)

    For amide hydrolysis, as mentioned above, acylation can be rate-limiting (k2<<k3). Then equation 7 becomes:

    v = k2EoS/[Ks+ S]

    Vm = k2Eo and Km = Ks

    Just as we saw before for the rapid equilibrium assumption (when ES falls apart to E + S more quickly than it goes to product), Km = Ks in amide hydrolysis.

    1. changing the pH or ionic strength - which can give data about general acids/bases in the active site:
    • a graph of kcat as a function of pH indicates that a group of pKa of approx. 6 must be deprotonated to express activity (i.e. Vm/2 is at about pH 6). This suggests that an active site histidine is necessary, which if it must be deprotonated to express activity, must be acting as a general base.
    • a graph of kcat/Km shows a bell-shaped curve indicating the necessity of a deprotonated side chain with a pKa of about 6 (i.e. the same His above) and a group which must be protonated with a pKa of about 10. This turns out to be an N terminal Ile (actually at the 16 position in the inactive precursor of chymotrypsin called chymotrypsinogen, which on activation of chymotrypsinogen loses the first 15 amino acids by selective proteolysis), which must be protonated to form a stabilizing salt bridge in the protein.

    Note: A new theoretical computer program, called THEMATICS (theoretical microscopic titration curves) has been developed to calculate the titration curves for all ionizable groups in a protein. When performed on test proteins, those amino acids that showed anomalous curves (flattened compared to normal titration curves) where usually found in the active site of the protein. The flattened curves show that the amino acid is partially protonated over a wider range of pH then theoretically expected. The program can be used to predict active site regions on protein of known structure but unknown function, and will be useful in the emerging field of proteomics. (figure below from Proc. Natl. Acad. Sci. USA, Vol. 98, Issue 22, 12473-12478, October 23, 2001)

    Fig. 1. Sample theoretical titration curves. Predicted mean net charge as a function of pH. (A) All of the histidine residues in the A chain of TIM: His-26 (+), His-95 (�), His-100 (*), His-115 (open square), His-185 (filled square), His-195 (open circle), His-224 (filled circle), and His-248 (). (B) Selected tyrosine residues of AR: Tyr-39 (+), Tyr-48 (�), Tyr-177 (*), Tyr-291 (open square), and Tyr-309 (filled square). (C) Selected lysine residues of PMI: Lys-100 (+), Lys-117 (�), Lys-128 (*), Lys-136 (open square), and Lys-153 (filled square). TIM, triosephosphate isomerase; AR, aldose reductase; PMI, phosphomannose isomerase. Remember that depending on the protein microenvironment, the pKa of a side chains like Asp can vary from 0.5 to 9.2!

    1. the enzyme - by chemical modification of specific amino acids, or through site-specific mutagenesis:
    1. modification of chyrmotrypsin (and many other proteases) with diisopropylphosphofluoridate (DIPF) modifies one of many Ser residues (Ser 195), suggesting that it is hypernucleophilic. and probably the amino acid which attacks the carbonyl C in the substrate, forming the acyl-intermediate.

    Figure: diisopropylphosphofluoridate

    Malathion and ethyl parathion (organic phosphate pesticides) have similar structures and reactivities as DIPF, and selectively inhibit serine proteases in insects.

    1. modification of the enzyme with tos-L-Phe-chloromethyl ketone inactives the enzyme with a 1:1 stoichiometry which results in a modified His.

    Figure: tos-L-Phe-chloromethyl ketone

    1. comparison of the primary sequence of many proteases show that three residues are invariant: Ser 195, His 57, Asp 102.
    2. site-specific mutagenesis show that if Ser 195 is changed to Ala 195, the enzymatic activity is almost reduced to background.

    Figure: Serine Protease Mechanism

    • The deprotonated His 57 acts as a general base to abstract a proton from Ser 195, enhancing its nucleophilicty as it attacks the electrophilic C of the amide or ester link, creating the oxyanion tetrahedral intermediate. Asp 102 acts electrostatically to stabilize the positive charge on the His.
    • The oxyanion collapses back to form a double bond between the O and the original carbonyl C, with the amine product as the leaving group. The protonated His 57 acts as a general acid donating a proton to the amine leaving group, regenerating the unprotonated His 57.
    • The mechanism repeats itself only now with water as the nucleophile, which attacks the acyl-enzyme intermediate, to form the tetrahedral intermediate.
    • The intermediate collapses again, releasing the E-SerO- as the leaving group which gets reprotonated by His 57, regenerating both His 57 and Ser 195 in the normal protonation state. The enzyme is now ready for another catalytic round of activity.
    • The mechanism for the first nucleophilic attack (by Ser) is the same as for the second (by water). The reverse mechanism of condensation of two peptide would be the reverse of the above mechanism, and is an example of the principle of microscopic reversibility.

    In short, all the catalytic mechanisms we encountered previously are at play in chymotrypsin catalysis. These include nucleophilic catalysis (with the Ser 195 forming a covalent intermediate with the substrates), general acid/base catalysis with His 57, and loosely, electrostatic catalysis with Asp 102 stabilizing not the transition state or intermediate, but the protonated form of His 57. An important point to note is that His, as a general acid and base catalyst, not only stabilizes developing charges in the transition state, but also provides a path for proton transfer, without which reactions would have difficulty in proceeding. One final mechanism is at work. The enzyme does indeed bind the transition state more tightly than the substrate. Crystal structures with poor "pseudo"-substrates that get trapped as partial tetrahedrally-distorted substrates of the enzyme and with inhibitors show that the oxyanion intermediate, and hence presumably the TS, can form H-bonds with the amide H (from the main chain) of Gly 193 and Ser 195. These can not be made to the trigonal, sp2 hybridized substrate. In the enzyme alone, the hole into which the oxyanion intermediate and TS would be placed is not occupied. This oxyanion hole is occupied in the tetrahedral intermediate.

    Figure: oxyanion hole in serine proteases: TS stabilzation

    Web Links:

    Serine Protease Home Page

    Jmol: Updated Chymotrypsin:D-Leu-L-Phe-p-fluorobenzylamde complex Jmol14 (Java) | JSMol (HTML5)

    Jmol: Updated Chymotrypin-Phenylethylboronate Inhibitor Complex Jmol14 (Java) | JSMol (HTML5)

    Many enzymes have active site serines which act as nucleophilic catalysts in nucleophilic substitution reactions (usually hydrolysis). One such enzyme is acetylcholine esterase which cleaves the neurotransmitter acetylcholine in the synapse of the neuromuscular junction. The transmitter leads to muscle contraction when it binds its receptor on the muscle cell surface. The transmitter must not reside too long in the synapse, otherwise muscle contraction will continue in an uncontrolled fashion. To prevent this, a hydrolytic enzyme, acetylcholine esterase, a serine esterase found in the synapse, cleaves the transmitter, at rates close to diffusion controlled. Diisopropylphosphofluoridate (DIPF) also inhibits this enzyme which effectively makes it a potent chemical warfare agent. An even more fluoride-based inhibitor of this enzyme, sarin, is the most potent lethal chemical agent of this class known. Only 1 mg is necessary to kill a human being.

    Figure: sarin


    Proteases Mechanisms

    Serine proteases are just one type of endoproteases. However, they are extremely abundant in both prokaryotes and eukaryotes. Protease A, a chymotrypin-like protease from Stremptomyces griseus, has a very different primary sequence than chymotrypsin, but its overall tertiary structure is quite similar to chymotrypsin, The positions of the catalytic triad amino acids in the primary sequences of the protein are very similar, indicating that the genes for the proteins diverged from a common precursor gene. In contrast, subtilisin, a serine protease from B. Subtilis, has both limited sequence and tertiary structure homology to chymotrypsin. However, when folded it also has a catalytic triad (Ser 221 - His 64 - Asp 32) similar to that of chymotrypsin (Ser 195 - His 57 - Asp 102).

    Jmol: Updated Comparison of Chymotrypin and Subtilisin Jmol14 (Java) | JSMol (HTML5)

    • Serine Protease Home Page: Structure Analyzer.

    Proteases have multiple functions, other than in digestion, including degrading old or misfolded proteins and activating precursor proteins (such as clotting proteases and proteases involved in programmed cell death). In general, four different classes of proteases have been found, based on residues found in their active sites. Proteases can also be integral membrane proteins, and carry out their activities in the hydrophobic environment of the membrane. For example, aberrant cleavage of the amyloid precursor protein by the membrane protease presenillin can lead to the development of Alzheimers.

    Figure: amyloid precursor protein

    Classification of Proteases

    Class (active site) Active Site Nucleophile Location Examples
    Serine/Threonine Hydrolases Ser/Thr soluble trypsin, chymotrypsin, subtilisin, elastase, clotting enzymes, proteasome
    membrane Rhomboid family
    Aspartic Hydrolases H2O activated by 2 Asps soluble pepsin, cathepsin, renin, HIV protease
    membrane β-secretase (BACE), presenilin I, signal peptide peptidase
    Cysteinyl Hydrolases Cys soluble bromelain, papain, cathespsins, caspases
    membrane ?
    Metallo Hydrolases H2O activated by 1 or 2 metal ions soluble thermolysin, angiotensin converting enzyme
    membrane S2P family
    Glutamate Hydrolases Glu . eqolysins (fungal)
    Asparagine Lysases (EC4) (elimination rx which are self-cleavage and hence not catalytic) Asn . Tsh autotransporter E. Coli

    β-secretase (BACE) is a membrane protein that contains two necessary Asp residues in its ectodomain (extracellular domain) which are used in the first cleavage of the N terminal domain of the beta amyloid precursor protein to release a soluble, N-terminal fragment of about 100,000 MW. γ-secretase, necessary for the second cleavage which frees the Aβ peptide is a heterotetramer composed of presenillin-1, nicastrin, APH-1 and PEN-2, and is located in neural plasma membranes and endoplasmic reticulum. The Aβ peptide moves to the extracellular side of the neural membrane where it aggregates. The remaining cytoplasmic part of the beta-amyloid precursor protein may regulate transcription. The presenillin subunit has protease activity. γ-secretase also cleaves another cell surface receptor protein, Notch. When this receptor has bound an extracelluar ligand, γ-secretase cleaves Notch within the cytoplasm, and the released fragment modifies gene transcription. The APH-1 subunit appears to inhibit presenilin protease activity while PEN-2 promotes it. Inhibiting γ-secretase would be an effect treatment for Alzheimers, but might have serious side effects since Notch processing would also be affected.

    Figure: Cleavage of beta amyloid precursor protein: protease and cofactors

    The γ-secretase protein quartet, and its roles in brain development and Alzheimer's disease. Presenilin-1, nicastrin, APH-1 and PEN-2 form a functional γ-secretase complex, located in the plasma membrane and endoplasmic reticulum (ER) of neurons. The complex cleaves Notch (left) to generate a fragment (NICD) that moves to the nucleus and regulates the expression of genes involved in brain development and adult neuronal plasticity. The complex also helps in generating the amyloid β-peptide (Aβ; centre). This involves an initial cleavage of the amyloid precursor protein (APP) by an enzyme called BACE (or β-secretase). The γ-secretase then liberates Aβ, as well as an APP cytoplasmic fragment, which may move to the nucleus and regulate gene expression. Mutations in presenilin-1 that cause early-onset Alzheimer's disease enhance γ-secretase activity and Aβ production, and also perturb the ER calcium balance. Consequent neuronal degeneration may result from membrane-associated oxidative stress, induced by aggregating forms of Aβ (which create Aβ plaques), and by the perturbed calcium balance. Reprinted by permission from Nature: Mattson, M. Nature. 422, 385-387 (2003)

    How do integral membrane protease catalyze the hydrolysis (using water) of transmembrane domains in proteins, given the hydrophobic environment of the bilayer? The rhomboid class of membrane proteases, which are found in prokaryotic and eukaryotic cells, is one of the most conserved membrane proteins in nature. Sequence comparison of rhomboid proteins shows a possible catalytic serine residue that is buried to the same depth in the bilayer as the cleavage site for rhomboid protein substrates. In addition the proteases share conserved amino acids similar to the catalytic triad of soluble serine proteases, with the critical amino acid localized on different membrane-spanning regions of the protein. The membrane triad amino acids are Ser, His and Asn (instead of Asp as in classical serine proteases). Classical serine protease inhibitors prevent cleavage of membrane protein substrates, but given the difficulty of reconstituting a functional purified rhomboid protein in vitro, the identity of specific amino acids in the catalytic mechanism is inferential at present.

    Helical wheel diagrams of putative transmembrane domains of the rhomboids (determined through hydropathy plots) show several to be amphiphilic. This might allow water, necessary for hydrolysis, to enter into the catalytic active site of the enzyme, although as was noted before, water has a reasonably high permeability through a membrane bilayer. The chief requirement for protein substrates of rhomboids is the presence of a transmembrane domain in the target protein. No specific amino acid sequence seems to be required for specificity of one particular substrate, the drosophila transmembrane protein spitz found in Golgi membranes. On cleavage of this protein, the remaining part of the protein is released as a water soluble protein to the lumen of the Golgi where it can eventually be released from the cell. The soluble protein fragment that is released from the cell contains an epidermal growth factor domain.

    Recently the structure of a rhomboid protease, GlpG, from E. Coli, was determined. This transmembrane protein has 6 transmembrane helices. The enzyme has a polar active site at the bottom of a V-shape opening situated laterally in the membrane. Active site His and Asn residue and many water molecules are deep in this V-shaped cleft well below the surface of the membrane. Access to the transmembrane strand of the protein substrate is blocked by a loop, which must be gated open to allow substrate access between the V-shaped gap between helices S1 and S3. Ser 201 (nucleophile) and His 254 (general base/acid) are essential for activity.

    Jmol: Updated E. Coli GlpG rhomboid intramembrane protease Jmol14 (Java) | JSMol (HTML5)

    CRISPR-Cas 9


    The CRISPR (clustered regularly interspaced short palindromic repeats) operon was initially discovered as part of the adaptive immune system of bacteria and archea, which must defend themselves against viruses (bacteriophages) and unwanted plasmid transferred from both bacteria. It would be ideal for bacteria to recognize previous exposure to viruses and their nucleic acids as the basis of their immunological memory system. Given the tendency of viral DNA to integrate into the host genome (which allows later transcription and translations of the viral genes in the process of new virus production), immunological memory could be based on that viral integrated DNA. Without going into detail, viral DNA can be integrated between two direct repeats in the bacterial genome. DNA from different viruses from previous exposures is also incorporated in the same fashion. One site of integration is the CRISPR operon. The DNA of the CRISPR operon contains both protein coding and noncoding regions which are transcribed and processed to form at least three RNA molecules (see figure below):

    • a coding Cas 9 mRNA this is translated to produce the Cas 9 (CRISPR associated protein);
    • a noncoding cr-RNA (CRISPR RNA)
    • a noncoding tracr-RNA (trans-activating CRISPR RNA)

    CRISPR operon and transcipts

    The two mature noncoding RNAs eventually associate to form a binary complex. When using CRISPR-Cas 9 in eukaroytic gene editing applications, the two noncoding RNAs are covalently combined into one large synthetic guide RNA (sg-RNA), described later in this section. The Cas 9 protein is an endonuclease that cleaves both strands of bound target dsDNA in a blunt-end fashion at specific sequences . This occurs after the DNA binds to two arginines (1333 and 1335) in Cas9 through a short (3-5+ bases) recognition protospacer adjacent motif (PAM) located three base pairs from the cleavage site. The DNA must also bind in a complementary and specific fashion to the protein-bound noncoding cr+tracr-RNAs (or a single sg-RNA molecule for gene editing applications). Binding and cleavage of target DNA would obviously render a recognized DNA from an invading bacteriophage inactive.

    Basic research into the bacterial CRISPR system has led to revolutionary and explosive applications of this gene editing system in eukaroytes. The hope is that CRISPR technology will give us a precise and incredibly cheap way to do gene therapy in diseased cells and organisms.

    We have discussed the structure and function of many proteins. Protein enzymes are key to life as they catalyze almost all biological reactions. Most key enzymes are regulated. The activity of Cas 9 must be carefully controlled. Think of the consequences if the enzyme were to cleave promiscuously at off-site targets! This section will help you understand several critical features of this enzyme:

    1. How does the enzyme find its correct target site, a 20 nucleotide DNA sequence and a proximal PAM site, among all the possible alternative sites. Think of how many PAM sequences there must be in the host DNA genome!
    2. How can the enzyme be "turned" on when it finds its target site and remain off when free, but more importantly, when it is bound off-site?

    First we will discuss the apo- form of the enzyme without bound substrate and RNA.

    ApoCas 9

    This section will focus on the Type II-A Cas9 from Streptococcus pyogenes (SpyCas9 or SpCas9). Cas 9 is an endonuclease that cleaves both strands of DNA 3 base pairs from a DNA motif, NCC/NGG, called PAM. It has two distinct lobes. The nuclease lobe (NUC), amino acids 1-56 and 718-1368, has two different nuclease domains for the two cleavages. The recognition or receptor lobe (REC), amino acids 94-717, interacts with the RNA molecules. There is also an arginine-rich bridge helix (57-93).

    The enzyme has two catalytic nuclease domains:

    • HNH-like nuclease domain which cleaves the "target" DNA strand, which is complementary to the RNA the confers specificity to the enzyme. The key catalytic residues are His 840 and Asn 854. It also contains a Mg ion;
    • Ruv-like domain that cleaves the complementary "non-target" strand with key active site residues Asp 10, Glu 762, Asp 986 and His 983. It also contains a bound Mn ion. The two lobes are separated by two linkers, amino acids 712-717, and an arginine-rich bridge (basic helix - BH), amino acids 628-658.

    The overall structure of the apoenyzme (without bound RNA and DNA,pdb id 4cmp) is shown in the figure below, which shows the NUC domain (light blue) with the two catalytic domains (HNH and Ruv), the REC domain (orange) and the BH helix (red).


    A close up view showing the two catalytic sites is shown in the figure and Jsmol below.


    A Jsmol of both the apo and holo forms of the enzyme can be found at the link below.

    Jmol: Cas 9 (4cmp - apo and 4008 - holo) Jmol14 (Java) | JSMol (HTML5)

    A comparison of the crystal structure of the apo-Cas 9 and the ternary Cas 9: sgRNA:DNA target strand complex shows a significant conformational change on binding nucleic acids. The structure of the holoenzyme (ternary complex) is shown in the figure and in the Jsmol below.


    The extent of the conformation change between apo- and holo-Cas 9 enzymes can be seen by examining the distance between D435 and E 944/945 in the figure below. The importance of this change will be described later.

    combo CRISPR Surf Conf Change

    The figure below shows the pathway from transcription of the relevant CRISPR genes (coding and noncoding) to the assembly of the ternary complex and the blunt end cut of the target DNA strand three nucleotides from the PAM sequence.


    The image below shows an expanded view of the ternary complex.


    Mechanism of DNA binding and cleavage

    The above figures do not speak to the mechanism of the binding processes that form the ternary complex. Kinetic and structural studies have been conducted to elucidate the mechanism of binding and cleavage and address the following questions:

    • which binds first, the RNA or DNA?
    • What are the consequences of the profound conformational changes on formation of the ternary complex?

    The specificity of target DNA binding depends both on enzyme:PAM DNA and enzyme:sgRNA (or tracr- and crRNA) interactions. It should seem improbable that the trinucleotide PAM DNA sequence (NGG in S. pyogenes), which interacts with a pair of arginines (R 1333, R 1335) through H-bonding, as shown in the images above, and other local sites in Cas 9 would provide the sole or even the majority of the binding interactions. The figure below shows the Args:PAM interaction (pdb code 4un3)


    Hence it is most likely that RNA binds first. Indeed, it does with the tracRNA implicated in recruitment of Cas and the crRNA providing specificity for target DNA binding. The resulting Cas9:RNA binary complex could then search the relevant DNA genome. That would include the DNA of the bacteriophage in viral infection or eukaryotic DNA if the CRISPR DNA operon with the genes for Cas 9 and a sg-RNA was transfected into the eukaryotic cell. After RNA binding, the enzyme would change conformation and allow loose DNA binding through Cas 9: PAM interactions.

    Studies have shown that the apo form can also bind DNA, but it does so loosely and indiscriminately. It dissociates quickly and binding is affected by generic polyanions such as the glycosoaminoglycan heparin, which indicates its nonspecific nature. Once bound, both off-target and target DNAs would then be surveyed. If a target DNA contained a PAM sequence, the complex would undergo another conformational change to position the HNH and Ruv nuclease catalytic residues and locally unwind the duplex DNA to make the blunt-end cuts.

    Cas 9 binding to the PAM site would promote better interaction of the unwound DNA and the bound RNA. If no PAM was present, no catalytically-effective Cas 9:target DNA would form. This prevents off-site cleavage. These allosteric changes and controls are vital to the function of the endonuclease. Here are some findings that support this proposed mechanism:

    • the conformation of apo Cas 9 is catalytically inactive;
    • on binding RNA to form a binary complex, Cas 9 undergoes a dramatic conformational change, mostly in the REC lobe. However on binding DNA in a nonspecific fashion, the conformational changes are much smaller. This suggests that most changes in conformation occur before DNA binding. In a way, RNA acts as a allosteric activator of the enzyme (as well as the major source of binding specificity to target DNA). Conformational changes can be determined directly by comparison of crystal structures or spectral techniques such as fluorescence resonance energy transfer (FRET) between two different attached fluorophores.
    • Cas 9: RNA interactions lead to ordering of the region of the RNA that interacts with the DNA PAM sequence and adjacent deoxynucleotides (a "seed sequence"), allowing the Cas 9:RNA complex to scan and interact with potential DNA targets with PAM sequences;
    • Once a PAM site is found, conformational changes leads to unwinding of the dsDNA, which allows heteroduplex formation between the crRNA and the target DNA strand;
    • since Cas 9 recognizes a variety of DNA target sequences (but of course only a specific PAM sequence), the binding of the target sequence depends on the geometry, not the sequence, of the target DNA;
    • since binding of off-target DNA to the Cas 9:RNA complex occurs but with very infrequent cleavage, binding and cleavage are very distinct steps;
    • on specific DNA binding, the HNH catalytic site moves near to the sessile DNA bond site. Crystal structures shown that the active site His is not sufficiently close to facilitate cleavage, suggesting that binding of a second metal ion (see below) may be necessary. Molecular dynamics studies show that the HNH domain is "remarkably plastic".

    The animated image below shows the relative conformational changes going from the apo Cas 9 to the binary Cas 9:sgRNA complex to the ternary Cas 9: sgRNA: target DNA complex. The NUC catalytic domain is shown in light blue, the REC (receptor or RNA binding domain) in orange, , sgRNA in red, and target DNA in green. Note again that on binding RNA to form a binary complex, Cas 9 undergoes a dramatic conformational change, mostly in the REC lobe. The pdb protein sequences shown were aligned using pdbEfold.

    animated image Cas 9

    A potential abbreviated catalytic mechanism for the Ruv nuclease domain is shown in the figure below. The red arrows indicate the second set of electron movements. His 983 acts as a general base to abstract a proton from water making it a more potent nucleophile. An intermediate trigonal bipyramidal phospho-intermediate is formed, which, along with the preceding transition state, is stabilized by the proximal Mg2+ ion (an example of electrostatic or metal ion catalysis). The magnesium is positioned through its interaction with negatively charged carboxyl groups of Asp 10, Glu 762, and Asp 986.


    More recent data suggests a second metal ion might be recruited to the Ruv site to further facilitate cleavage of the DNA. The HNH catalytic site has a structure (beta-beta-alpha) and conserved His in common with a class of nucleases that require one metal ion. In contrast, the Ruv catalytic site does not have this common secondary structural motif and has a critical histidine, both common features found in endonucleases that use two metal ions.

    CRISPR and Eukaryotic Gene Editing

    How could blunt-end cutting of both strands of DNA by Cas 9 lead to the holy grail of specific eukaryotic gene editing with no off-site effects? Cutting the DNA genome seems like a bad idea. It fact, it is potentially so bad that a myriad of DNA repair mechanisms have evolved to fix the cut. These include homologous recombination. If corrective DNA is supplied as well as the components of the CRISPR system, a cell could effectively add the corrective DNA after the double-stranded cut and repair a deleterious mutation. Consult a molecular biology textbook for more insight into homologous recombination.

    Mutations in the PAM sequence prevent Cas9 nuclease activity. Hence the NGG PAM sequence is vital for the interactions and activity described above. This would seem to limit the utility of CRISPR-Cas 9 in eukaryotic gene editing, until one realizes that the GG dinucleotide has a 5.2% frequency of occurrence in the human genome, which corresponds to over 160 million occurrences. Even then it might not occur in a desire gene target. Cas 9 nuclease from other bacteria extend the range of activity of the CRISPR/Cas system as they interact with other PAM sequences (NNAGAA and NGGNG for S. thermophilus and NGGNG for N. meningidtis). Likewise, mutations in the S. pyogenes PAM (NGG) have been made as well. A D1135E mutation retains but increases the specificity for the normal NGG PAM site. D1135V, R1335Q and T1337R mutations alter the optimal PAM recognition site to NGAN or NGNG.

    CRISPR editing can be easily used to knock out specific genes. In addition, if cells are transfected with a plasmid with many target sequences, the system can be used to edit multiple genes in one experiment. This would be very useful in studies of diseases linked to multiple genes. Since the cost of CRISPR reagents (plasmids, RNAs) is so inexpensive, and the specificity of editing is so high, the great excitement about CRISPR use for gene editing in human disease and for modification of plant and fungal genomes is warranted.

    Other systems have been developed to specifically bind to a target DNA sequence and then cleave it. They typically contain a protein that binds to a specific DNA target and an associated endonuclease that cleaves within the target DNA site. Typical prokaryotic restriction enzymes bind to and cut at a specific nucleotide sequence (for example Eco R1 cleaves at G/AATTC palindromic sequences) to form sticky ends. The protein itself binds to this DNA recognition site. Other examples are based on the structure of known transcription factors. Libraries of genetically engineered proteins with Zn finger DNA binding domains (designed for specific DNA target sequences) fused to endonucleases have been created for this purpose. Another example are proteins called TALENs (transcription activator-like effector nucleases). These are fusion proteins containing a TAL effector DNA-binding domain and a nuclease. In each of these cases, a 3D-folded protein is the specific target DNA recognition molecule. Think how much easier it is to make in effect a 1D-DNA recognition element, a simple linear RNA sequence, which would adopt the correct 3D structure on binding of its complementary target.

    One major problem in the use of CRISPR for gene editing must be solved: how to get the CRISPR components in the correct cells in an organism. In effect, it's the same problem faced by small drug designers only the components are much larger. Ex vivo applications, when diseased cells are removed from the body, repaired by CRISPR, and then reinjected, are likely to have more success. In these cases, electroporation would allow uptake of Cas 9 and the sg-RNA. In vivo therapy has included use of adeno-associated viruses in which genes for Cas 9 and sg RNA could be encapsulated. This technique, used for other gene delivery systems, has the advantage of being tolerated immunologically. However, this system allows for continual gene expression which is undesirable for gene editing. After an initial "fix" of a mutant gene, continued expression of the CRISPR-Cas 9 genes would increase the chances for off-target cutting. A more recent approach is to delivery the mRNA in artificial lipid nanoparticles that can be taken into cells. Once free and translated into protein and sg RNA inside the cell, gene editing has a chance to occur before the RNA and protein are degraded.

    Other Information we may want to include in this section:

    Overview of Catalytic Mechanisms

    The descriptions below describe the major mechanisms enzymes use to catalyze chemical reactions. It should be noted that many enzymes use more than one, and sometimes several different catalytic strategies during their reaction mechanism.

    Covalent Catalysis

    Covalent catalysis involves the formation of a covalent bond between the enzyme and at least one of the substrates involved in the reaction. Often times this involves nucleophilic catalysis which is a subclass of covalent catalysis. As seen in Section 7.1, several amino acid R-groups can serve as a nucleophile and are often found at the active site of enzymes. Nucleophilic side chains are often activated by deprotonation caused by neighboring side chains, such as histidine that can act as a base. Alternatively, water can also activate the nucleophile. The intermediate covalent bond formation between the enzyme and the substrate enables bond cleavage and the removal of a leaving group.

    Acid-Base Catalysis

    Acid-Base Catalysis is involved in any reaction mechanism that requires the transfer of a proton from one molecule to another. It is very common to see this mechanism combined with Covalent Catalysis as many nucleophiles are activated by the removal of a proton, including alcohol, thiol, and amine functional groups. Enzymes that utilize Acid-Base Catalysis can be subgrouped further into either specific acid-base or general acid-base reactions. Specific acid or specific base catalysis occur if a hydronium ion (H3O+) or a hydroxide ion (OH-), respectively, are utilized directly in the reaction mechanism, and the pH of the solution affects the rate of catalysis. General acid and general base reactions occur when molecules other than hydronium ion (H3O+) or a hydroxide ion (OH-) are the source of proton donation or acceptance. Most commonly, an active site amino acid residue is used to accept or donate a proton within the reaction mechanism. In general acid-base reactions the pH is usually held constant within a buffered system.

    Electrostatic Catalysis

    Electrostatic catalysis occurs when the enzyme active site stabilizes the transition state of the reaction by forming electrostatic interactions with the substrate. The electrostatic interactions can be ionic, ionic-dipole, dipole-dipole, or hydrophobic interactions. Hydrogen bonding is one of the most common electrostatic interactions formed in the active site.


    Enzyme active sites can become devoid of water and mimic the reaction characteristics of the gas phase. This can destabilize the polarized state of charged groups such as acids and bases. Thus, the neutral form of these types of residues becomes the favored state. This is due to significant alterations in the pKa of the active site residues within the nonpolar environment. This can cause normally acidic residues such as glutamate to abstract a proton from histidine and behave as a base, for example.

    Catalysis by Approximation

    In catalysis by approximation, the enzyme enhances the reaction rate by binding with multiple substrates and positioning them favorably so that the reaction can proceed. Binding with the enzyme reduces the rotational entropy of the substrates that would otherwise be randomly free floating in solution, and enables the correct positioning of substrates for the reaction. The loss of entropy, which is not favorable, is offset by the binding energy released with the substrate-enzyme interaction.

    In addition to correctly positioning the substrates to interact with one another, catalysis by approximation converts a reaction that would have been second order, with substrates that are free floating in solution, to a first order reaction, where all of the substrates are held in place by the enzyme and behave as a single molecule. This can dramatically improve the catalytic rate of the reaction from 105 to 107 times faster, depending on the enzyme system.

    Strain Distortion

    In organic chemistry, you learned that certain structures such as three-membered and four-membered ring structures, such as epoxides were highly reactive due to the strain distortion inherent to the unfavored bond angles inherent to the ring. Enzyme active sites can also utilize strain distortion within a bound substrate to increase the reactivity of the molecule and favor the formation of the transition state. Many enzymes that function by the induced fit model also utilize strain distortion within their catalytic mechanism. Within the unbound state they remain in a low catalytic state, however the interaction with the substrate induces the destabilization of the enzyme active site or may induce strain within the substrate causing the initiation of the catalytic activity of the enzyme.

    Cofactor Catalysis

    Cofactors are molecules that bind to enzymes and are required for the catalytic activity of the enzyme. They can be divided into two major categories: metals and coenzymes. Metal cofactors that are commonly found in human enzymes include: iron, magnesium, manganese, cobalt, copper, zinc, and molybdenum. Coenzymes are small organic molecules that are often derived from vitamins, which are essential organic nutrients consumed within the diet. Coenzymes can bind loosely with the enzyme and have the ability to bind and release from the active site, or they may be tight binding and lack the ability to release easily from the enzyme. Tight binding coenzymes are referred to as prosthetic groups. Enzymes that are not yet associated with a required cofactor are called apoenzymes, whereas enzymes that are bound with their required cofactors are called holoenzymes. Sometimes organic molecules and metals combine to form coenzymes, such as in the case of the heme cofactor (Figure 7.15). Coordination of heme cofactors with their enzyme counterparts often involves electrostatic interactions with histidine residues as shown in the succinate dehydrogenase enzyme shown in Figure 7.15.

    Figure 7.15 The Heme Cofactor. The family of heme cofactors contain an iron metal coordinated with a porphyrin ring structure as shown in the left hand panel within the structure of Heme B. In the right hand panel, Heme B is shown complexed with the succinate dehydrogenase enzyme from the Kreb Cycle.

    Structure of Heme B shown in the left hand panel is from: Yikrazuul and the crystal structure of Succinate Dehydrogenase complexed with Heme B is from: Richard Wheeler.

    As an example of how vitamins can be utilized as cofactors, Table 7.2 shows the common B vitamins and the coenzymes derived from their structures. Many vitamin deficiencies cause disease states due to the inactivity of apoenzymes that are unable to function without the correctly bound coenzyme.

    Table 7.2 Essential B-Vitamins and their Modified Enzyme Cofactors

    Cofactors can help to mediate enzymatic reactions through the use of any of the different catalytic strategies listed above. They can serve as nucleophiles and mediate covalent catalysis or form electrostatic interactions with the substrate and stabilize the transition state. They can also cause strain distortion or facilitate acid-base catalysis. Metal-aided catalysis can often use homolytic reaction mechanisms that involve radical intermediates. This can be important in reactions such as those occurring in the electron transport chain that require the safe movement of single electrons.

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    7.3 Examples of Reaction Mechanisms

    Protease Enzymes

    Proteolytic enzymes (also termed peptidases, proteases and proteinases) are capable of hydrolyzing peptide bonds in proteins. They can be found in all living organisms, from viruses to animals and humans. Proteolytic enzymes have great medical and pharmaceutical importance due to their key role in biological processes and in the life-cycle of many pathogens. Proteases are extensively applied enzymes in several sectors of industry and biotechnology, furthermore, numerous research applications require their use, including production of Klenow fragments, peptide synthesis, digestion of unwanted proteins during nucleic acid purification, cell culturing and tissue dissociation, preparation of recombinant antibody fragments for research, diagnostics and therapy, and the exploration of the structure-function relationships.

    Proteolytic enzymes belong to the hydrolase class of enzymes and are grouped into the subclass of the peptide hydrolases or peptidases. Depending on the site of enzyme action the proteases can also be subdivided into exopeptidases or endopeptidases. Exopeptidases, such as aminopeptidases and carboxypeptidases catalyze the hydrolysis of the peptide bonds near the N- or C-terminal ends of the substrate, respectively (Figure 7.16). Endopeptidases (Figure 7.16) cleave peptide bonds at internal locations within the peptide sequence. Proteases may also be nonspecific and cleave all peptide bonds equally or they may be highly sequence specific and only cleave peptides after certain residues or within specific localized sequences.

    Figure 7.16 Common Peptidase Reactions. Aminopeptidases (top diagram) and carboxypeptidases (middle diagram) remove the terminal amino acid residues, whereas endopeptidases (lower diagram) cleave protein sequences at internal sites. The red arrows show the peptide bonds to be cleaved.

    Figure from: Mótyán, J.A., et al. (2013) Biomolecules 3(4), 923-942

    The action of proteolytic enzymes is essential in many physiological processes. For example, proteases function in the digestion of food proteins, protein turnover, cell division, the blood-clotting cascade, signal transduction, processing of polypeptide hormones, apoptosis and the life-cycle of several disease-causing organisms including the replication of retroviruses such as the human immunodeficiency virus (HIV). Due to their key role in the life-cycle of many hosts and pathogens they have great medical, pharmaceutical, and academic importance.

    It was estimated previously that about 2% of the human genes encode proteolytic enzymes and due to their necessity in many biological processes, proteases have become important therapeutic targets. They are intensively studied to explore their structure-function relationships, to investigate their interactions with the substrates and inhibitors, to develop therapeutic agents for antiviral therapies or to improve their thermostability, efficiency and to change their specificity by protein engineering for industrial or therapeutic purposes.

    Based on the catalytic mechanism and the presence of amino acid residue(s) at the active site the proteases can be grouped as aspartic proteases, cysteine proteases, glutamic proteases, metalloproteases, asparagine proteases, serine proteases, threonine proteases, and proteases with mixed or unknown catalytic mechanism. Here we will explore the reaction mechanism and sequence specificity of the family of serine proteases.

    The list of serine proteases is quite long. They are grouped in two broad categories - 1) those that are chymotrypsin-like and 2) those that are subtilisin-like. Though subtilisin-type and chymotrypsin-like enzymes use the same mechanism of action, including the catalytic triad, the enzymes are otherwise not related to each other by sequence and appear to have evolved independently (Figure 7.17). They are, thus, an example of convergent evolution - a process where evolution of different forms converge on a structure to provide a common function.

    Figure 7.17 Convergent Evolution of the Serine Proteases, Chymotrypsin and Subtilisin. The crystal structure of the eukaryotic, Bovine Chymotrypsin (left hand panel) with catalytic triad indicated in green. The crystal structure of the prokaryotic Subtilisin BPN from the bacterium Bacillus subtilis (right hand panel) with the catalytic triad and common mutations indicated using the ball and stick models. Note that the catalytic triad formation is strikingly similar between the two structures, whereas the surrounding protein structures and sequence do not show homology or related ancestry.

    Image of bovine chymotrypsin modified from Mattyjenjen and image of subtilisin BNP from Romero-Garcia, E.R., et al. (2009) J Biomed Biotech 2009(1):201075

    The chymotrypsin-like serine protease enzymes cleave the peptide bond on the carboxylic acid side of specific amino acids and the specificity is determined by the size/shape/charge of amino acid side chain that fits into the enzyme’s S1 binding pocket (Figure 7.18). Three chymotrypsin-like family members that share high sequence homology are the pancreatic digestive enzymes, trypsin, chymotrypsin and elastase. The protein cleavage sites of these enzymes varies. Trypsin cleaves proteins on the carboxylic side of basic residues, such as lysine and arginine, while Chymotrypsin cleaves after aromatic hydrophobic amino acids, such as phenylalanine, tyrosine, and tryptophan, and Elastase cleaves after small, hydrophobic residues, such as glycine, alanine, and valine. As shown in Figure 7.18, variations in the amino acid residues within the binding pocket of these proteases, enables electrostatic interactions with the substrate and determines sequence specificity.

    Figure 7.18 Substrate Specificity of Trypsin, Chymotrypsin, and Elastase. The upper panel shows the space-filling crystal structures of Trypsin, Chymotrypsin, and Elastase, respectively, with the S1 substrate binding pocket indicated. The lower panel depicts the S1 binding domains of each protease in more detail with important amino acid R-groups indicated. For Trypsin, an aspartate residue in the lower portion of the S1 pocket aid in electrostatic interactions with basic residues of the substrate. The Chymotrypsin S1 binding pocket is large and hydrophobic in nature accommodating aromatic residues of the substrate, while the Elastase S1 binding pocket is small and hydrophobic, only allowing other small and hydrophobic R-groups to dock in this location.

    Image modified from: Goodsell, D. (2012) Molecule of the Month, Protein Database and Aleia Kim

    Serine proteases use four of the major catalytic mechanism during the reaction cycle: Acid-Base Catalysis, Covalent Catalysis, Electrostatic Interactions, and Desolvation. The active site of serine proteases contains a catalytic triad of three amino acids: His, Ser (hence the name "serine protease") and Asp. These three key amino acids each play an essential role in the cleaving ability of the proteases. While the amino acid members of the triad are located far from one another in the primary sequence of the protein, due to folding, they will be very close to one another in the heart of the enzyme.

    During a catalysis event, an ordered mechanism occurs in which several intermediates are generated. The catalysis of the peptide cleavage can be seen as a ping-pong catalysis, in which a substrate binds (in this case, the polypeptide being cleaved), a product is released (the N-terminus "half" of the peptide), another substrate binds (in this case, water), and another product is released (the C-terminus "half" of the peptide). Figure 7.19 details the catalytic process.

    In step 1 to 2, the polypeptide substrate enters the active site and is positioned in the correct orientation near the active site serine residue through electrostatic interactions in the S1 binding pocket. The carbonyl carbon of the substrate is positioned near the active site serine residue. The hydrogen of the serine alcohol is abstracted by the catalytic histidine residue through acid-base catalysis. This is made possible by the action of the active site aspartate residue. In this case, the aspartate residue abstracts a proton from histidine, enabling histidine to remove the proton from the serine alcohol. Normally, this would be an odd thing for aspartate to be able to do, as the pKa of the R-group of aspartate is much lower than that of histidine within an aqueous environment. However, when the peptide substrate docks in the active site of the enzyme, this excludes water from the area through a desolvation process, and creates a hydrophobic microenvironment. This effectively raises the pKa of aspartate and favors the uncharged or protonated form of the residue, causing the proton abstraction from histidine.

    In steps 2 to 3 covalent catalysis is enabled as the active site serine mediates nucleophilic attack on the carbonyl carbon of the substrate forming a tetrahedral oxyanion intermediate. The oxyanion intermediate in the pathway is then stabilized by electrostatic interactions with a region of the protease known as the oxyanion hole. Here the negative charge of the oxyanion is stabilized by electrostatic interactions with amide nitrogens from the protease backbone. Rebounding of the oxyanion to reform the carbonyl double bond leads to the cleavage of the peptide bond. The C-terminal portion of the protein can then leave the active site of the enzyme.

    Once the C-terminal peptide leaves the active site, water can enter and rehydrate the active site, as seen in steps 5 to 6. A water molecule is oriented in proximity to the carbonyl carbon of the N-terminal peptide that is still bound to the active site serine residue. The oxygen from water acts as a nucleophile and attacks the carbonyl carbon, recreating an oxyanion intermediate which is stabilized by the electrostatic interactions of the oxyanion hole. As this oxyanion rebounds to reform the carbonyl, the serine residue acts as a leaving group and the N-terminal peptide is released from the enzyme. The presence of water in the active site, re-establishes the serine residue and the native states of the histidine and aspartic acid residues.

    Overall, each amino acid in the triad performs a specific task in this process:

    • The serine has an -OH group that is able to act as a nucleophile, attacking the carbonyl carbon of the scissile peptide bond of the substrate (covalent catalysis).
    • A pair of electrons on the histidine nitrogen has the ability to accept the hydrogen from the serine -OH group, thus coordinating the attack of the peptide bond (acid/base catalysis).
    • The carboxyl group on the aspartic acid in turn coordinates with the histidine, making the nitrogen atom mentioned above much more electronegative through the process of desolvation.

    Electrostatic interactions are critical to (1) bind the substrate in the S1 binding pocket (2) stabilize the transition state oxyanion, and (3) coordinate the water molecule to mediate nucleophilic attack on the enzyme-bound intermediate.

    Figure 7.19 Reaction mechanism of Chymotrypsin-like Proteases. The reaction mechanism has been broken down into an eight step process. In steps 1-3 the protein substrate binds with the protease and is oriented to place the carbonyl carbon of the substrate in proximity with the active site serine residue. Acid-base catalysis enables the activation of the serine residue to mediate nucleophilic attack on the protein substrate. The covalent oxyanion intermediate shown in 3 and 4 is stabilized by the oxyanion hole. Rebound of the electrons to reform the carbonyl group cause the cleavage of the peptide bond and the removal of the C-terminal portion of the peptide from the active site, shown in 5. Water enters the active site and mediates nucleophilic attack on the carbonyl carbon of the enzyme-substrate intermediate, as shown in 6 and 7. As the carbonyl bond is reformed, serine acts as a leaving group and the N-terminal peptide is released from the enzyme. The catalytic triad in the enzyme active site is recovered and the enzyme is reset for another round of catalytic activity, as shown in 8.

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    Adenylate Kinases

    Adenylate kinase (also known as AK or myokinase) is a phosphotransferase enzyme that catalyzes the interconversion of adenine nucleotides (ATP, ADP, and AMP). By constantly monitoring phosphate nucleotide levels inside the cell, AK enzymes play an important role in cellular energy homeostasis. The basic chemical reaction mediated by this enzyme class is the conversion of 2 ADP molecules into 1 ATP and 1 AMP (Figure 7.20A). The reverse reaction can also occur forming an equilibrium based on cellular concentrations of the varying phosphorylation states.

    To date there have been nine human AK protein isoforms identified. While some of these are ubiquitous throughout the body, some are localized into specific tissues. For example, AK7 and AK8 are both only found in the cytosol of cells; and AK7 is found in skeletal muscle whereas AK8 is not. Not only do the locations of the various isoforms within the cell vary, but the binding of substrate to the enzyme and kinetics of the phosphoryl transfer are different as well. AK1, the most abundant cytosolic AK isozyme, has a Km about a thousand times higher than the Km of AK7 and 8, indicating a much weaker binding of AK1 to AMP. Sub-cellular localization of the AK enzymes is done by unique targeting sequences found in the protein. Each isoform also has different preference for NTP's. Some will only use ATP, whereas others will accept GTP, UTP, and CTP as the phosphoryl carrier.

    AK enzymes can be involved in regulating nucleotide concentrations and serve as a relay system between cellular and mitochondrial pools of adenine nucleotides, as shown in Figure 7.20B. AK enzymes can also serve as a sensor for energy load within the cell and can lead to the activation of AMP-sensitive systems within the cell when energy levels are low (Figure 7.20C).

    Figure 7.20. Adenylate Kinase (AK) Enzyme Activity. (A) The fundamental reaction of AMP Kinases. (B) reactions of AK enzymes can work in cascading mechanisms to provide signaling and communication between different regions of the cell, including the cytoplasm and mitochondria, and (C) AK enzymes are often used as a metabolic monitor of energy load, leading to the activation or inhibition of downstream enzymes.

    Figure modified from: Dzeja, P. and Terzic, A. (2009). Int J Mol Sci 10(4):1729-1772

    Phosphoryl transfer during the AK reaction only occurs after the closing of an 'open lid' structure in the enzyme through the catalysis by approximation mechanism (Figures 7.21 and 7.22). This causes an exclusion of water molecules that brings the substrates in proximity to each other and effectively lowers the energy barrier for the nucleophilic attack by the γ-phosphoryl group of ATP on the α-phosphoryl of AMP. In the crystal structure of the AK enzyme from E. coli with inhibitor Ap5A (Figure 7.21C), the Arg88 residue coordinate the Ap5A at the α-phosphate group through electrostatic interactions. It has been shown that the mutation of Arg88 to Gly (R88G) results in 99% loss of catalytic activity of this enzyme, suggesting that this residue is intimately involved in the phosphoryl transfer. Another highly conserved residue is Arg119, which lies in the adenosine binding region of the AK, and acts to sandwich the adenine in the active site. It has been suggested that the promiscuity of these enzymes in accepting other NTP's is due to this relatively inconsequential interactions of the base in the ATP binding pocket. A network of positive, conserved residues (Lys13, Arg123, Arg156, and Arg167 in AK from E. coli) stabilize the buildup of negative charge on phosphoryl group during the transfer. Two distal aspartate residues bind to the arginine network, causing the enzyme to fold and reduces its flexibility. A magnesium cofactor is also required, essential for increasing the electrophilicity of the phosphate on AMP, though this magnesium ion is only held in the active pocket by electrostatic interactions and dissociates easily.

    Flexibility and plasticity allow proteins to bind to ligands, form oligomers, aggregate, and perform mechanical work. Large conformational changes in proteins play an important role in cellular signaling. AK acts as a signal transducing protein; thus, the balance between conformations regulates protein activity. AK has an 'open' conformation (Figure 7.21A) that is induced into the 'closed' and biologically active conformation upon substrate binding.

    Figure 7.21 Crystal Structure of the Adenylate Kinase Enzyme. Structures of the open (A, PDB ID: 4AKE) and the closed (B, PDB ID: 1AKE) states. The LID and the NMP domains are shown in red and orange respectively. The CORE domain and the rest of the protein are shown in blue. (C) PDB image 3HPQ showing the AK enzyme skeleton in cartoon and the key residues as sticks and labeled according to their placement in the E. coli AK enzyme, crystallized with Ap5A inhibitor.

    Figures A and B modified from: Das, A., et al. (2014) PLoS Computational Biology 10(4):e1003521 and Figure C from: Snodgrah

    Within the AK protein structure, there is a core domain and two smaller domains called the LID and NMP (Figure 7.22). ATP binds in the pocket formed by the LID and CORE domains. AMP binds in the pocket formed by the NMP and CORE domains. Localized regions of the protein fold and unfold during conformational transitions in the reaction mechanism and enhance the catalytic efficiency.

    The two subdomains (LID and NMP) can fold and unfold independently of one another depending on substrate binding. Substrate binding induces regional shifts in the protein structure to the partially closed or fully closed conformations. The fully closed conformation optimizes alignment of substrates for phosphoryl-transfer and aids with the removal of water from the active site to avoid wasteful hydrolysis of ATP.

    Figure 7.22 Conformational Transition pathway and proposed catalytic mechanism of AK. Model a, substrate free AK with an open conformation. Model b, ATP bound form of ADK with a closed LID domain. Model c, ATP and AMP bound form of AK with a closed conformation. Model d, two ADP bound forms of AK with a closed conformation. Model e, one ADP bound form of AK with a closed NMP domain.

    Figure from: Ping, J., et al, (2013) BioMed Res Int: 628536

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    Restriction Endonucleases

    A restriction enzyme, restriction endonuclease, or restrictase is an enzyme that cleaves DNA into fragments at or near specific recognition sites within molecules known as restriction sites. Restriction enzymes are one class of the broader endonuclease group of enzymes. Restriction enzymes are commonly classified into five types, which differ in their structure and whether they cut their DNA substrate at their recognition site, or if the recognition and cleavage sites are separate from one another. To cut DNA, all restriction enzymes make two incisions, once through each sugar-phosphate backbone (i.e. each strand) of the DNA double helix. Here we will focus on the Type II restriction enzymes that are routinely used in molecular biology and biotechnology applications.

    As with other classes of restriction enzymes, Type II Restriction Enzymes occur exclusively in unicellular microbial life forms––mainly bacteria and archaea (prokaryotes)––and are thought to function primarily to protect these cells from viruses and other infectious DNA molecules. Inside a prokaryote, the restriction enzymes selectively cut up foreign DNA in a process called restriction digestion; meanwhile, host DNA is protected by a modification enzyme (a methyltransferase) that modifies the prokaryotic DNA and blocks cleavage. Together, these two processes form the restriction modification system.

    The first Type II Restriction Enzyme discovered was HindII from the bacterium Haemophilus influenzae Rd. The event was described by Hamilton Smith (Figure 7.23) in his Nobel lecture, delivered on 8 December 1978:

    ‘In one such experiment we happened to use labeled DNA from phage P22, a bacterial virus I had worked with for several years before coming to Hopkins. To our surprise, we could not recover the foreign DNA from the cells. With Meselson’s recent report in our minds, we immediately suspected that it might be undergoing restriction, and our experience with viscometry told us that this would be a good assay for such an activity. The following day, two viscometers were set up, one containing P22 DNA and the other Haemophilus DNA. Cell extract was added to each and we began quickly taking measurements. As the experiment progressed, we became increasingly excited as the viscosity of the Haemophilus DNA held steady while the P22 DNA viscosity fell. We were confident that we had discovered a new and highly active restriction enzyme. Furthermore, it appeared to require only Mg2+ as a cofactor, suggesting that it would prove to be a simpler enzyme than that from E. coli K or B.

    After several false starts and many tedious hours with our laborious, but sensitive viscometer assay, Wilcox and I succeeded in obtaining a purified preparation of the restriction enzyme. We next used sucrose gradient centrifugation to show that the purified enzyme selectively degraded duplex, but not single-stranded, P22 DNA to fragments averaging around 100 bp in length, while Haemophilus DNA present in the same reaction mixture was untouched. No free nucleotides were released during the reaction, nor could we detect any nicks in the DNA products. Thus, the enzyme was clearly an endonuclease that produced double-strand breaks and was specific for foreign DNA. Since the final (limit) digestion products of foreign DNA remained large, it seemed to us that cleavage must be site-specific. This proved to be case and we were able to demonstrate it directly by sequencing the termini of the cleavage fragments.’

    Figure 7.23. Hamilton Smith and Daniel Nathans at the Nobel Prize press conference, 12 October 1978 (reproduced with permission from Susie Fitzhugh). Original Repository: Alan Mason Chesney Medical Archives, Daniel Nathans Collection.

    Image from: Pingoud, A., Wilson, G.G., and Wende, W. (2014) Nuc Acids Res 42(12):7489-7527.

    Restriction enzymes are named according to the taxonomy of the organism in which they were discovered. The first letter of the enzyme refers to the genus of the organism and the second and third to the species. This is followed by letters and/or numbers identifying the isolate. Roman numerals are used to specify different enzymes from the same organism. For example, the enzyme ‘HindIII’ was discovered in Haemophilus influenzae, serotype d, and is distinct from the HindI and HindII endonucleases also present within this bacterium. The DNA-methyltransferases (MTases) that accompany restriction enzymes are named in the same way, and given the prefix ‘M.’. When there is more than one MTase, they are prefixed ‘M1.’, ‘M2.’, etc, if they are separate proteins or ‘M1∼M2.’ when they are joined.

    Restriction Enzymes that recognize the same DNA sequence, regardless of where they cut, are termed ‘isoschizomers’ (iso = equal; skhizo = split). Isoschizomers that cut the same sequence at different positions are further termed ‘neoschizomers’ (neo = new). Isoschizomers that cut at the same position are frequently, but not always, evolutionarily drifted versions of the same enzyme (e.g. BamHI and OkrAI). Neoschizomers, on the other hand, are often evolutionarily unrelated enzymes (e.g.EcoRII and MvaI).

    Type II Restriction Enzymes are a conglomeration of many different proteins that, by definition, have the common ability to cleave duplex DNA at a fixed position within, or close to, their recognition sequence. This cleavage generates reproducible DNA fragments, and predictable gel electrophoresis patterns, properties that have made these enzymes invaluable reagents for laboratory DNA manipulation and investigation. Almost all Type II Restriction Enzymes require divalent cations, usually Mg2+, as essential components of their catalytic sites. Ca2+, on the other hand, often acts as an inhibitor of Type II Restriction Enzymes.

    The recognition sequences of Type II Restriction Enzymes are palindromic in nature, with two possible types of palindromic sequences. The mirror-like palindrome is similar to those found in ordinary text, in which a sequence reads the same forward and backward on a single strand of DNA, as in GTAATG. The inverted repeat palindrome is also a sequence that reads the same forward and backward, but the forward and backward sequences are found in complementary DNA strands (i.e., of double-stranded DNA), as in GTATAC (GTATAC being complementary to CATATG). Inverted repeat palindromes are more common and have greater biological importance than mirror-like palindromes. The position of cleavage within the palindromic sequence can vary depending on the enzyme and can produce either single stranded overhanging sequences (sticky ends) or blunt-ended DNA products.

    The EcoRI restriction enzyme produces sticky ends:

    EcoRI restriction enzyme recognition site.svg

    whereas SmaI restriction enzyme cleavage produces blunt-ends:

    SmaI restriction enzyme recognition site.svg

    Methylation can be used by the host to protect its own genome from cleavage. For example, the methylation of the EcoRI recognition sequence by the M.EcoRI methyltransferase (MTase), changes the sequence from GAATTC to GAm6ATTC (m6A = N6-methyladenine). This modification completely protects the sequence from cleavage by EcoRI.

    Type II Restriction Enzymes initially bind non-specifically with the DNA and proceed to slide down the DNA scanning for recognition sequences (Fig 7.24). Upon binding to the correct palindromic sequence the enzyme associates with the metal cofactor and mediates catalytic cleavage of the DNA using the mechanism of strain distortion and catalysis by approximation.

    Figure 7.24 DNA Recognition and Cleavage by Type II Restriction Endonucleases. (A) Pictorial view of an EcoRV dimer scanning nonspecifically along the DNA until a specific binding site is recognized. This causes coupling with the metal cofactor and strain distortion of the DNA. Hydrolysis of the phosphodiester bond is mediated and the DNA cleavage products released from the enzyme. (B) shows a space filling model of EcoRV DNA recognition and cleavage.

    Figure (A) from: Pingoud, A., Wilson, G.G., and Wende, W. (2014) Nuc Acids Res 42(12):7489-7527. and Figure (B) from: Thomas Splettstoesser

    One of the most important questions regarding the catalytic mechanism of a hydrolase is whether hydrolysis involves a covalent intermediate, as is typical for the proteases described previously. This can be decided by analyzing the stereochemical course of the reaction. This was done first for EcoRI, and later for EcoRV. Both enzymes were found to cleave the phosphodiester bond with inversion of stereoconfiguration at the phosphorus, which argues against the formation of a covalent enzyme–DNA intermediate. Thus, it is proposed that cleavage involves the direct nucleophilic attack of the substrate by a water molecule, as shown in Figure 7.25.

    Figure 7.25 A General Mechanism for DNA Cleavage by EcoRI and EcoRV. An activated water molecule attacks the phosphorous in-line wiht the phosphodiester bond to be cleaved, which proceeds with inversion of configuration. X, Y, and Z are a general base, a Lewis acid and a general acid, respectively.

    Figure from: Pingoud, A., Wilson, G.G., and Wende, W. (2014) Nuc Acids Res 42(12):7489-7527.

    Type II restriction enzymes typically form a homodimer when binding with DNA, as shown in the crystal structure of BglII in Figure 7.26B. BglII catalyses phosphodiester bond cleavage at the DNA backbone through a phosphoryl transfer to water. Studies on the mechanism of restriction enzymes have revealed several general features that seem to be true in almost all cases, although the actual mechanism for each enzyme is most likely some variation of this general mechanism (Figure 7.25). This mechanism requires a base to generate the hydroxide ion from water, which will act as the nucleophile and attack the phosphorus in the phosphodiester bond. Also required is a Lewis acid to stabilize the extra negative charge of the pentacoordinated transition state phosphorus, as well as a general acid or metal ion that stabilizes the leaving group (3’-O). In some Type II Restriction Enzymes, two divalent metal cofactors are required (such as in EcoRV and BamHI), whereas other enzymes only require one divalent metal cofactor (such as in EcoRI and BglII).

    Structural studies of endonucleases have revealed a similar architecture for the active site with the residues following the weak consensus sequence Glu/Asp-(X)9-20-Glu/Asp/Ser-X-Lys/Glu. BglII's active site is similar to other endonucleases', following the sequence Asp-(X)9-Glu-X-Gln. In its active site there sits a divalent metal cation, most likely Mg2+, that interacts with Asp-84, Val-94, a phosphoryl oxygen, and three water molecules. One of these water molecules, is able act as a nucleophile because of its proximity to the scissile phosphoryl group (Figure 7.26A). The nucleophilic water molecule is positioned for attack onto the phosphoryl group by a hydrogen bond with the side chain amide oxygen of Gln-95 and its contact with the metal cation. Interaction with the metal cation effectively lowers its pKa, promoting the water's nucleophilicity (Figure 7.26 A). During hydrolysis, the divalent cation is able to stabilize the 3'-O- leaving group and coordinate proton abstraction from one of the coordinated water molecules (Figure 7.26A).

    Figure 7.26 Proposed Reaction Mechanism for the Type II Restriction Endonuclease, BglII. (A) Schematic diagram of the catalytic mechanism demonstrating the utility of Mg2+ ions and polar amino acid residues within the active site to activate and position a water molecule for nucleophilic attack on the phosphodiester bond of the DNA substrate. (B) Crystal structure of the BglII dimer with double stranded DNA and (C) Coordination of the Mg2+ cofactor within the active site of the BglII enzyme.

    Figures from: GWilliams

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    Ribozymes (ribonucleic acid enzymes) are RNA molecules that have the ability to catalyze specific biochemical reactions, including RNA splicing in gene expression, similar to the action of protein enzymes. In 1982, the self-splicing Group I intron was reported as the first discovered catalytic RNA. It was described in the ciliate protozoa Tetrahymena thermofila by Sidney Altman and Thomas Czech, who were awarded the Nobel Prize in Chemistry in 1989. Similar introns can be found in some prokaryotic genomes and the mitochondria and chloroplast DNA of diverse eukaryotes. The second example of a ribozyme to be discovered was the RNAse P involved in tRNA maturation, which had a key biological role and a ubiquitous occurrence in both prokaryotes and eukaryotes. The third reported catalytic RNA was a tiny ribozyme (~50 nt), the self-cleaving hammerhead ribozyme (HHR), which was found in a group of atypical plant pathogens with small circular RNA (circRNA) genomes such as viral satellite RNAs and viroids. Since then, a few more examples of either natural or artificial ribozymes have been discovered, including the ribosome, a singular catalytic RNA that catalyzes the peptide bond formation, the central chemical reaction in extant biology. This landscape strongly supports the hypothesis of a prebiotic RNA world, where the first self-replicating organisms were based on RNA as both the genetic material and as catalyst. Whereas modern proteins would have replaced most of these ancient catalytic RNAs, some of them have remained in current organisms performing different functions. Among all known ribozymes, there is the enigmatic family of small (<200 nt) self-cleaving RNAs, which catalyse a simple intramolecular transesterification in a highly sequence-specific manner. This reaction, which can occur spontaneously in the RNA, starts by a SN2-like nucleophilic attack of the 2′-oxygen to the adjacent 3′-phosphate, resulting in cleavage of the phosphodiester bond to form a 2′-3′-cyclic phosphate and a 5′-hydroxyl RNA products (Figure 7.27A).

    Figure 7.27 The Hammerhead Ribozyme. (A) Mechanism of internal transesterification reaction in the RNA. The cleavage reaction proceeds with an attack of the hydroxyl moiety at 2′ to the phosphate group at 3′, followed by a bipyramidal transition-state. The cleavage products are a 2′-3′-cyclic phosphate at the 5′ RNA product and a 5′-hydroxyl at the 3′ RNA product; (B) Diagram of the hammerhead ribozyme. Black boxes indicate the highly conserved nucleotides at the catalytic core. Secondary structures of (C) 3-Dimensional structure of the Hammerhead Ribozyme.

    Figures A and B from: De la Peña, M., et al (2017) Molecules 22(1):78 and Figure C from: Wgscott

    Similar to ribonuclease proteins such as the RNAse A, small self-cleaving ribozymes stabilize the formation of the bipyramidal oxyphosphorane transition-state through different catalytic strategies, such as in-line atomic orientation, electrostatic neutralization, and general acid-base catalysis. In this way, nucleolytic ribozymes are able to catalyze RNA cleavage at a rate only a few-fold slower than their protein counterparts. At least nine classes of naturally-occurring small self-cleaving ribozymes have been described so far: the hammerhead (Figure 7.27C), hairpin, human Hepatitis-δ, Varkud-satellite, GlmS, twister, twister sister, hatchet and pistol ribozymes. Since its discovery 30 years ago, the HHR has been extensively used as a model ribozyme for structural, biochemical and biological studies. It is composed of a catalytic centre comprising 15 highly conserved nucleotides surrounded by three double helixes (I to III), which adopt a secondary structure that resembles the shape of a hammerhead shark head. Depending on the open-ended helix, there are three possible circularly permuted forms, named type I, II or III (Figure 7.27B). The HHR motif, like other small ribozymes such as hairpin and Hepatitis-δ, has been historically regarded as a biological peculiarity of subviral circular RNA genomes. However, we know now that small catalytic RNAs such as the HHR can occur numerously in DNA genomes from bacteria to eukaryotes, including our own genome, and carry out diverse biological functions that we are just starting to recognize.

    The ribosome also functions as a ribozyme mediating the formation of the amide bind during protein synthesis. Protein synthesis from a mRNA template occurs on a ribosome, a nanomachine composed of proteins and ribosomal RNAs (rRNA). The ribosome is composed of two very large structural units that are an amalgamation of proteins and rRNA molecules (Figure 7.28). The smaller unit (termed 30S and 40S in bacteria and eukaryotes, respectively) coordinates the correct base pairing of the triplet codon on the mRNA with another small adapter RNA, transfer or tRNA, that brings a covalently connected amino acid to the site.

    Figure 7.28 The Small Ribosomal Subunit of Thermus thermophilus. The 16S ribosomal RNA is shown in orange with ribosomal proteins attached in blue.

    Figure from: Goodsell, D.S. - Molecule of the Month

    Peptide bond formation occurs when another tRNA-amino acid molecule binds to an adjacent codon on mRNA. The tRNA has a cloverleaf tertiary structure with some intrastranded H-bonded secondary structure. The last three nucleotides at the 3' end of the tRNA are CpCpA. The amino acid is esterified to the terminal 3'OH of the terminal A by a protein enzyme, aminoacyl-tRNA synthetase.

    Covalent amide bond formation between the second amino acid to the first, forming a dipeptide, occurs at the peptidyl transferase center, located on the larger ribosomal subunit (50S and 60S in bacteria and eukaryotes, respectively). The ribosome ratchets down the mRNA so the dipeptide-tRNA is now at the the P or Peptide site, awaiting a new tRNA-amino acid at the A or Amino site. Figure 7.29 shows a schematic of the ribosome with bound mRNA on the 30S subunit and tRNAs covalently attached to amino acid (or the growing peptide) at the A and P site, respectively.

    Figure 7.29 Schematic Representation of a Bacterial Ribosome. The 50s (yellow) and 30s (blue) subunits of the ribosome are composed of protein and rRNA. mRNA (red linear strand) is shown docked onto the 30s subunit. The P and A sites are filled with tRNA molecules (green and red).

    Figure from: Jakubowski, H.

    A likely mechanism for the formation of the amide bond between a growing peptide on the P-site tRNA and the amino acid on the A-site tRNA has been derived from crystal structures with bound substrates and transition state analogs and is shown in Figure 7.30. Catalysis does not involve any of the ribosomal proteins (not shown) since none is close enough to the peptidyl transferase center to provide amino acids that could participate in general acid/base catalysis. Hence the rRNA must acts as the enzyme (i.e. it is a ribozyme). Initially it was thought that a proximal adenosine with a perturbed pKa could, at physiological pH, be protonated/deprotonated and hence act as a general acid/base in the reaction. However, none was found. The most likely mechanism to stabilize the oxyanion transition state at the electrophilic carbon attack site is precisely located water, which is positioned at the oxyanion hole by H-bonds to uracil 2584 on the rRNA. The cleavage mechanism involves the concerted proton shuffle shown Figure 7.30. In this mechanism, the substrate (Peptide-tRNA) assists its own cleavage in that the 2'OH is in position to initiate the protein shuttle mechanism. (A similar mechanism might occur to facilitate hydrolysis of the fully elongated protein from the P-site tRNA.) Of course all of this requires perfect positioning of the substrates and isn't that what enzymes do best? The main mechanisms for catalysis of peptide bond formation by the ribosome (as a ribozyme) are intramolecular catalysis and transition state stabilization by the appropriately positioned water molecule.

    Figure 7.30 Mechanism of Peptide Bond Formation. Peptide bond formation is likely mediated by a proton shuttle mechanism that is stabilized by a coordinated water molecule within the oxyanion hole.

    Figure from: Jakubowski, H.

    The crystal structure of the eukaryotic ribosome has recently been published (Ben-Shem et al). It is significantly larger (40%) than the prokaryotic counterpart, with mass of around 3x106 Daltons. The 40S subunit has one rRNA chain (18S) and 33 associated proteins, while the larger 60S subunit has 3 rRNA chains (25S, 5.8S and 5S) and 46 associated proteins. The larger size of the eukaryotic ribosome facilitates more interactions with cellular proteins and greater regulation of cellular events. The structure of a eukaryotic 80S ribosome showing rRNA and protein interactions is shown in Figure 7.31.

    Figure 7.31 Eukaryotic 80S Ribosome. The 40S subunit is on the left, the 60S subunit on the right. The ribosomal RNA (rRNA) core is represented as a grey tube, expansion segments are shown in red. Universally conserved proteins are shown in blue. These proteins have homologs in eukaryotes, archaea and bacteria. Proteins shared only between eukaryotes and archaea are shown in orange, and proteins specific to eukaryotes are shown in red. PDB identifiers 4a17, 4A19, 2XZM aligned to 3U5B, 3U5C, 3U5D, 3U5E

    Figure from: Fvoigtsh

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