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6.5: 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 kineticists, medicinal chemists and molecular biologists (i.e. a well trained chemist) can change:

    1. the substrate - for example, changing the leaving group or acyl substituents 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 - an odd idea on the surface but it leads to new insights into enzye catalysis.

    We will explore in detail the mechanisms of selected enzymes. For some we will concentrate mostly on reaction mechanisms based on structural data. For others, we will use kinetic and structural data to hypothesize a reaction mechanism consistent with all of the data. Even with lots of data, there are often different proposed mechanisms for a given reaction. Kinetic data is vital in that it can help to determine:

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


    This enzyme cleaves C-terminal amino acid from a protein through a hydrolysis reaction. As such it is an exoprotease, not an endoprotease which cleaves proteins internally within the sequence. In terms of selectivity toward C-terminal amino acids, its activity is increase if the C-terminal side chain group is aromatic or branched aliphatic (Phe, Tyr, Trp, Leu or Ile). 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+ might have several roles. In one, it may help a coordinated water to be more nucleophilic by either polarizing the water or converting it a the more potent nucleophile OH-. It might also stabilize developing negative charges in the transition state and in an intermediate. Two possible mechanisms have been offered.

    The Water pathway. In this reaction, water acts as a nucleophile, 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. A simplified reaction reaction is shown in Figure \(\PageIndex{1}\) below.


    Figure \(\PageIndex{1}\): Water pathway mechanism for carboxypeptide A. After Wu et al. J Phys Chem B. 2010 July 22; 114(28): 9259–9267. doi:10.1021/jp101448j

    Nucleophilic Pathway. In this pathway, Gu 270 is the primary initial nucleophile in the formation of the initial tetrahedral intermediate. The role of Zn2+ is in charge stabilization. This pathway is illustrated in Figure \(\PageIndex{2}\) below.

    Figure \(\PageIndex{2}\): Nucleophilic Pathway after Wu et al. J Phys Chem B. 2010 July 22; 114(28): 9259–9267. doi:10.1021/jp101448j

    Figure \(\PageIndex{3}\) below shows an interactive iCn3D model of the active site of bovine carbonic anhydrase in the absence of a substrate or inhibitor (1M4L). The Zn2+ ion is shown as a red sphere.

    Bovine carboxypeptidase A (1M4L).png

    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{3}\): Bovine carboxypeptidase A (1M4L) (Copyright; author via source).
    Click the image for a popup or use this external link:

    Note how far tyrosine 248 is away from the active site in the model. Glu72 and Glu270 are negatively charged in the resting state of the enzyme at pH 7.5. The values are much higher (weaker acid) than solution pka of the side chain of glutamic acid. Also the water bound to the Zn2+ is long enough to suggest that the water is neutral and not in the form of OH- in this form of the enzyme. If OH- were present, the distance between it and the Zn2+ would be shorter due to the great electrostatic force.

    Figure \(\PageIndex{4}\) below shows an interactive iCn3D model of the active site of bovine carbonic anhydrase bound to the inhibitor aminocarbonylphenylalanine (1HDU). The Zn2+ ion is shown as a red sphere.

    Bovine carboxypeptidase A inhibitor aminocarbonylphenylalanine (1HDU).png

    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{4}\): Bovine carboxypeptidase A bound to bound to the inhibitor aminocarbonylphenylalanine (1HDU). (Copyright; author via source).
    Click the image for a popup or use this external link:

    Note the closer proximity of tyrosine 248 to the active site.


    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 (NAG)6 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. This 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. A reaction mechanism of hemiacetal/acetal formation and cleavage is illustrated in Figure \(\PageIndex{5}\) below.

    Figure \(\PageIndex{5}\): A reaction mechanism of hemiacetal/acetal formation and cleavage

    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. Here is a possible general mechanism:

    • 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 stabilized carbocation, forming the hemiacetal with release of the extra proton from water to the deprotonated Glu 35 reforming the general acid catalysis.

    Part of a mechanism illustrating the roles of Glu 35 and Asp 52 is shown below in Figure \(\PageIndex{6}\).

    Figure \(\PageIndex{6}\): Mechanism of acetal cleavage by lysozyme

    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 catalyst and forms a covalent bond with NAM, expelling a NAG leaving group with Glu 35 acting as a general acid as shown in Figure \(\PageIndex{7}\) below. This alternative mechanism also is consistent with other β-glycosidic bond cleavage enzyme. Substrate distortion is also important in this alternative mechanism.

    Figure \(\PageIndex{7}\): Alternative mechanism for lysozyme catalysis employing Asp 52 as a nucleophilc catalyst

    Recent structural work shows that Asp 52 is involved in a strong hydrogen bond network that might preclude its ability to form a covalent bond with the glycan substrate. An earlier structure (1H6M) did show a covalent bond.

    Figure \(\PageIndex{8}\) below shows an interactive iCn3D model of the active site of hen egg white lysozyme bound to a (NAG)4 glycan (7BR5). Note the positions of E35 and D52.


    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{8}\): Interactions of hen egg white lysozyme with bound (NAG)4 glycan (7BR5) (Copyright; author via source).
    Click the image for a popup or use this external link:

    Chymotrypsin and othe endoproteases

    Chymotrypsin, an endoprotease, cleaves by hydrolysis an internal peptide bond after aromatic side chains. It also cleaves small ester and amide substrates after aromatic residues. It has a similar mechanism to a multitude of other proteases that used the same catalytic triad, Ser 195, Asp 102 and His 95, so we'll study it significant detail.

    It's easier in the lab to study the enzyme using small substrate mimics of a protein that two use a substrate protein. The mimics include both esters and amides. In determining the mechanism of an enzyme, you have to change an experimental variable and see how catalytic activity changes. What can be changed? Turns out everything including the solvent! Let's explore these changes and how they affect chymotrypsin activity.

    1. changing the substrate (for example changing the leaving group or acyl substituents of a hydrolyzable substrate):

    Data from the cleavage of small amide and ester substrates shown in Figure \(\PageIndex{9}\) suggest that a covalent intermediate occurs on chymotrypsin catalyzed cleavage.

    Figure \(\PageIndex{9}\): Small amide and ester substrates of chymotrypsin

    Table \(\PageIndex{1}\): below shows kinetic data for the cleavage of these substrates.

    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

    Here's how these data can be interpreted.

    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. 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 previously consistent with these ideas before. The data suggest a mechanism based on the chemcial equations shown in Figure \(\PageIndex{10}\) below:

    Figure \(\PageIndex{10}\): Chemical equations for chymotrypsin hydrolysis of a substrate involved a covalent intermediate with ping-pong kinetics.

    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. 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). The mathematical equation for this reaction is shown below (without derivation)

    \mathrm{v}_{0}=\frac{\left(\frac{\mathrm{k}_{2} \mathrm{k}_{3}}{\mathrm{k}_{2}+\mathrm{k}_{3}}\right) \mathrm{E}_{0} \mathrm{~S}}{\mathrm{~K}_{\mathrm{S}}\left(\frac{\mathrm{k}_{3}}{\mathrm{k}_{2}+\mathrm{k}_{3}}\right)+\mathrm{S}}

    For hydrolysis of ester substrates, which have better leaving groups compared to amides, deacylation is rate limiting, ( k3<<k2). For amide hydrolysis, as mentioned above, acylation can be rate-limiting (k2<<k3). From this, equation xx can be simplified as shown in Table \(\PageIndex{2}\) below for ester and amide hydrolysis.

    Ester hydrolysis (deacylation rate limiting, k3 << k2) Amide hydrolysis (deacylation rate limiting, k2 << k3)
    \mathrm{v}_{0}=\frac{\mathrm{k}_{3} \mathrm{E}_{0} \mathrm{~S}}{\mathrm{~K}_{\mathrm{S}}\left(\frac{\mathrm{k}_{3}}{\mathrm{k}_{2}}\right)+\mathrm{S}}
    \mathrm{v}_{0}=\frac{\mathrm{k}_{2} \mathrm{E}_{0} \mathrm{~S}}{\mathrm{~K}_{\mathrm{S}}+\mathrm{S}}
    V_{M}=k_{3} E_{0}
    \mathrm{V}_{\mathrm{M}}=\mathrm{k}_{2} \mathrm{E}_{0}

    Table \(\PageIndex{2}\):

    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 the case of 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: The PKAD is a database of experimentally measured pKa values of ionizable groups in proteins. It is searchable by the PDB ID.)

    1. change the enzyme - by chemical modification of specific amino acids, or through site-specific mutagenesis: Here are some specific examples.
    1. modification of chymotrypsin (and many other proteases) with diisopropylphosphofluoridate (DIPF) modifies only one (Ser 195) of many serines in the protein, suggesting that it is hypernucleophilic. and probably the amino acid which attacks the carbonyl C in the substrate, forming the acyl-intermediate. This reaction is illustrated in Figure \(\PageIndex{11}\) below. The figure also shows analagous molecules used in common insecticides, which act through a similar mechanism.
    Figure \(\PageIndex{1}\): Mechanism of inhibition of chymotrypin by covalent modification by diisopropylphosphflouridate.
    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, as shown in Figure \(\PageIndex{12}\) below.
    Figure \(\PageIndex{12}\): Reaction of chymotrypsin and other serine protease with chloromethy ketones.
    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. The strongly suggests that Ser 195 is an active site nucleophile.

    D. Changing the solvent. Yes indeed you can take chymotrypsin and show that it is active in anhydrous organic solvents. Surely this is impossible you say! It is true and we will explore it at the end of our discussion on proteases since its hard enough to understand chymotrypsin activity in aqueous solution. No new chemistry is needed, just a change in what your mind can conceptualize.

    Now, let's put all this together in the general mechanism for serine protease cleavage of protein, as shown in Figure \(\PageIndex{13}\) below.

    Figure \(\PageIndex{13}\): General mechanism of peptide bond cleavage by chymotrypsin and other serine proteases

    Here what the mechanism shows:

    • The deprotonated His 57 acts as a general base to abstract a proton from Ser 195, enhancing its nucleophilicity 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.

    A crystal structure of a relative of chymotrypsin, trypsin, which cleaves after positively charged lysine and arginine side chains has been determined with a bound transition state analog inhibitor. The transition state inhibitor is t-butoxy-Ala-Val-boro-Lys methyl ester shown in Figure \(\PageIndex{14}\) below.

    t-butoxy-Ala-Val-boro-Lys methyl ester.svg
    Figure \(\PageIndex{14}\): The transition state inhibitor of trypsin, t-butoxy-Ala-Val-boro-Lys methyl ester

    Everyone should remember from introductory chemistry that neutral boron compounds like BH3 and BF3 are trigonal planar (sp2) and electron deficient. Although the boron is not charged, it has a significant partial positive charge (δ+) so it is electrophilic. The nucleophilic oxygen of Ser 195 can then attack the boron to form a tetrahedral intermediate. This intermediate is not an oxyanion but one of the attached oxygens with a δ- charge occupies the oxyanion hole.

    Figure \(\PageIndex{15}\) below show the active site group in trypsin interacting with part of the transition state analog (1BTZ). The serine 195 side chain O is covalently attached to the boron, so the boron is now tetrahedral (sp3).

    Figure \(\PageIndex{15}\):

    The yellow dotted lines show hydrogen bonding between the backbone amide Hs of Ser 195 and Gly 193 with the methoxy oxygen of the now tetrahedral borate transition state inhibitor. The boron atom is the yellow/orange sp3 atom connected to 3 oxygen (red) atoms and one carbon (cyan) atom. Normally, the oxyanion O- from the tetrahedral intermediate in amide or ester cleavage would occupy the oxyanion hole.

    Figure \(\PageIndex{16}\) below shows an interactive iCn3D model of the active site of the phenylethane boronic acid (PBA) complex of alpha-chymotrypsin (6cha).


    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{16}\): Active site of the phenylethane boronic acid (PBA) complex of alpha-chymotrypsin (6cha). (Copyright; author via source).
    Click the image for a popup or use this external link:

    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 ( Figure \(\PageIndex{17}\)), is the most potent lethal chemical agent of this class known. Only 1 mg is necessary to kill a human being.

    Figure \(\PageIndex{17}\): Sarin

    Serine proteases have unique specificities to allow cleavage after a different subset of side chains. The 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 \(\PageIndex{18}\) below, variations in the amino acid residues within the binding pocket of these proteases, enables electrostatic interactions with the substrate and determines sequence specificity.

    Figure \(\PageIndex{18}\): 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

    A schematic nomenclature developed by Berger and Schechter is often used to show the sites on the substrate ( labeled P3, P2, P1, P1', P2' and P3') referring to the products made after cleavage of the peptide/protein that is cleaved between P1 and P' (the scissile bond) and the corresponding sites on the protease (S3, S2, S1, S1', S2' and S3'). This is illustrated in Figure \(\PageIndex{19}\) below.

    Figure \(\PageIndex{19}\): Nomenclature of denote key interacting groups in the enzyme (Sn) and in the substrate (Pn) around the scissile bond for protein cleavage

    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). The alignment of the core structures of chymotrypsin (5cha, magenta) and subtilisin (1sbc, cyan), are shown in Figure \(\PageIndex{20}\) below.

    Figure \(\PageIndex{20}\): Convergent Evolution of the Serine Proteases, Chymotrypsin and Subtilisin.

    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. They are, thus, an example of convergent evolution - a process where evolution of different forms converge on a structure to provide a common function.

    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.

    Table \(\PageIndex{3}\) below shows a classification of proteases based on their active site nucleophiles.

    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

    Table \(\PageIndex{3}\) below shows a classification of proteases based on their active site nucleophiles.

    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. Instead of using a catalytic triad, these serine proteases use a dyad of Ser 201 as a nucleophile and His 254 as a general acid/base..

    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.

    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. The active site His and Ser residues aare 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. The active site His 254 can be covalently modified with different chloromethylketone peptide derivatives. Figure \(\PageIndex{21}\) below shows an interactive iCn3D model of the Rhomboid intramembrane protease GlpG 4QO2

    Rhomboid intramembrane protease GlpG 4QO2.png

    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{21}\): Rhomboid intramembrane protease GlpG (4QO2). (Copyright; author via source).
    Click the image for a popup or use this external link:

    Proteolytic enzymes (also termed peptidases, proteases and proteinases) are 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 (like chymotrypsin) or endopeptidases (like carboxypeptidase A) as we have above Exopeptidases, such as aminopeptidases and carboxypeptidases catalyze the hydrolysis of the peptide bonds near the N- or C-terminal ends of the substrate, respectively. Endopeptidases cleave peptide bonds at internal locations within the peptide sequence. These differences are illustrated in Figure \(\PageIndex{22}\) below. 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 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.

    Enzyme catalysis in organic solvents

    We discussed changing the solvent and exploring its affect on enzyme catalysis. Seems crazy but let's see what happens. Attempts have been made to do this for the last 100 years. These including putting the enzyme in:

    • water miscible solvents like ethanol and acetone were added. If the water concentration was high enough, activity remained.
    • biphasic mixtures in which an aqueous solution of an enzyme was emulsified in a water immiscible solvent like chloroform or ethylacetate. The substrate would partition into both phases, while the product hopefully would end up into the organic phase.
    • nearly nonaqueous solvents, with a few % water at less than the solubility limits of water.
    • anhydrous organic solvents (0.01% water). It is this case that is most astonishing since enzymatic activity is often retained.

    It is important to realize that in this last case, the enzyme is not in solution. It is rather in suspension and acts as a heterogeneous catalyst, much like palladium acts as a heterogeneous catalyst in the hydrogenation of alkenes. The suspension must be mixed vigorously and then sonicated to produce small suspended particles, so diffusion of reactants into the enzyme and out is not rate limiting. Let's explore the activity of chymotrypsin in a nonpolar solvent. Consider the following questions.

    • Why aren't the enzymes inactive? Surely it must seem ridiculous that they aren't, since as we learned earlier, proteins are not that stable. A 100 amino acid protein on average is stabilized only about 10 kcal/mol over the denatured state, or the equivalent of a few H bonds. Surely the hydrophobic effect, one of the dominant contributors to protein folding and stability, would not stabilize the native structure of enzymes in nonpolar organic solvents, and the protein would denature. It doesn't however! Maybe the real question should be not whether water is necessary, but rather how much water is necessary. The enzyme can't "see" more than a monolayer or so of water around it. The data suggests that the nature of the organic solvent is very important. The most hydrophobic solvents are best in terms of their ability to maintain active enzymes! Chymotrypsin retains 104 more activity in octane than pyridine (see kcat/Km below), which is more hydrophilic than octane. The more polar the solvent, the more it can strip bound water away from the protein. If you add 1.5% water to acetone, the bound water increases from 1.2 to 2.4%, and the activity of chymotrypsin increases 1000 fold.

    Table \(\PageIndex{4}\) below shows chymotrypsin activity in organic solvents.

    Solvent Structure kcat/Km (M-1min-1) relative ratio
    H2O bound to enzyme (%, w/w)
    Octane tablec1.gif 63 15000x 2.5
    Toluene tablec2.gif 4.4 1000x 2.3
    Tetrahydrofuran tablec3.gif 0.27 175x 1.6
    Acetone tablec4.gif 0.022 5.5x 1.2
    Pyridine tablec5.gif <0.004 1x (.004) 1.0

    Table \(\PageIndex{4}\): Chymotrypsin activity in organic solvents

    • How active are enzymes in nonpolar solvents? Enzymes are often studied in model transesterification reactions. Typical reaction conditions are enzyme at 1 mg/ml, with one substrate, an ester such as N-acteyl-L-Phe-ethyl ester,at 2-12 mM, and the other substrate, an alcohol, such as n-propanol (instead of being water as in a typical hydrolysis reaction) at 0.25-1.5 M. The more concentrated alcohol replaces the alcohol (ethanol) esterified in the ester. Michaelis-Menten kinetics are followed, with biomolecular rate constants of 1010 greater than without the enzyme.
    • How much water do the enzymes need? 1 molecule of chymotrypsin in octane has less than 50 molecules of water associated and can demonstrate activity. To form a monolayer requires about 500 water molecules. Water can be added which presumably leads to more bound water and higher activity.
    • How stable are the enzymes? Denaturation requires conformational flexibility, which apparently requires water. The half-life of chymotrypsin in water at 60oC is minutes, but in octane at 100oC it is hours. At 20oC, the half-life in water is a few days, but in octane it is greater than 6 months. Remember two things contribute to stability. The protein can denature at high temperatures. Also since chymotrypsin is a protease, it can cleave itself in a autoproteolytic reaction.

    Table \(\PageIndex{5}\) below shows the half-life of chymotrypsin activity in water and octane

    Solvent 60oC 100oC 20oC
    water minutes - few days
    octane - hours > 6 months

    Table \(\PageIndex{5}\): Half-life of chymotrypsin activity in water and octane at different temperatures

    Is the enzyme specificity changed? The net binding energy is a function of the binding energy of the substrate - the binding energy of the water, since water must be displaced from the active site on binding. In an anhydrous solvent, specificity changes must be expected. For chymotyrpsin, the driving force for binding of substrates in water is mostly hydrophobic. In water, the kcat/Km for the reaction of N-acteyl-L-Ser-esters is reduced 50,000x compared to the Phe ester. However, in octane, chymotrypsin is three times more active toward Ser esters than Phe esters.

    Table \(\PageIndex{6}\) shows specificity changes in chymotrypsin in water and octane

    Substrate kcat/Km
    solvent: H2O solvent: Octane
    N-acetyl-L-Ser-ester 1x 3x
    N-acetyl-L-Phe-ester 50,000x 1x

    Table \(\PageIndex{6}\): Specificity changes in chymotrypsin in water and octane

    Now consider competitive inhibitors. Napthalene binds 18 times more tightly than 1-napthoic acid, but in octane, the chymotrypsin binds napthoic acid 310 times as tightly. Likewise the ratio of [kcat/Km (L isomer)]/[kcat/Km (D isomer)] of N-acetyl-D- or N-acetyl-L-Ala-chloroethyl esters is 1000-10,000 in water, but less than 10 in octane.

    Table \(\PageIndex{7}\) shows chymotrypsin inhibition constants in water and octane

    Inhibitor Inhibition Constant Ki (nM)
    In water In Octane
    Benzene 21 1000
    Benzoic acid 140 40
    Toluene 12 1200
    Phenylacetic acid 160 25
    Naphthalene 0.4 1100
    1-Naphthoic acid 7.2 3

    Table \(\PageIndex{7}\): Chymotrypsin inhibition constants in water and octane

    Can new reactions be carried out in nonpolar solvents? The quick answers is yes, since reactions in aqueous solutions can be unfavorable due to low Keq's, side reactions, or insolubility of reactants. Consider lipases which cleave fatty acid esters by hydrolysis in aqueous solutions. In nonaqueous solutions, reactions such as transesterification or ammonolysis can be performed.

    Enzymes are clearly active in organic solvents which appears to contradict our central concepts of protein stability. Two reasons could could explain this stability.

    1. It is possible that from a thermodynamic view, the enzyme is stable in organic solvents. However, as was discussed above, this is inconceivable given the delicate balance of noncovalent and hydrophobic interactions required for protein stability.
    2. The second reason must win the day: the protein is unable to unfold from a kinetic point of view. Conformational flexibility is required for denaturation. This must require water as the solvent. Denaturation in organic solvents is kinetically, not thermodynamically controlled.

    A specific example helps illustrate the effects of different solvents on chymotrypsin activity. Dry chymotrypsin can be dissolved in DMSO, a water miscible solvent. In this solvent it is completely and irreversibly denatured. If it is now diluted 50X with acetone with 3% water, no activity is observed. (In the final dilution, the concentrations of solvents are 98% acetone, 2.9% water, and 2% DMSO.) However, if dry chymotrypsin was added to a mixture of 98% acetone, 2.9% water, and 2% DMSO, the enzyme is very active. We end up with the same final solvent state, but in the first case the enzyme has no activity while in the second case it retains activity. These ideas are illustrated in Figure \(\PageIndex{23}\) below.

    Figure \(\PageIndex{23}\): Chymotrypsin activity in acetone depends on order of solvent addition

    Dry enzymes added to a concentrated water-miscible organic solvent (like DMSO) will dissolve and surely denature, but will retain activity when added to a concentrated water-immiscible solvent (like octane), in which the enzyme will not dissolve but stay in suspension.

    It appears the enzymes have very restricted conformational mobility in nonpolar solvents. By lyophilizing (freeze-drying) the enzyme against a specific ligand, a given conformation of a protein can be trapped or literally imprinted onto the enzyme. For example, if the enzyme is dialyzed against a competitive inhibitor (which can be extracted by the organic solvent), freeze-dried to remove water, and then added to a nonpolar solvent, the enzyme activity of the "imprinted" enzyme in nonpolar solvents is as much as 100x as great as when no inhibitor was present during the dialysis. If chymotrypsin is lyophilized from solutions of different pHs, the resulting curve of V/Km for ester hydrolysis in octane is bell-shaped with the initial rise in activity reaching half-maximum activity at a pH of around 6.0 and a fall in activity reaching half-maximum at pH of approximately 9.

    Use of enzymes in organic solvent allows new routes to organic synthesis. Enzymes, which are so useful in synthetic reactions, are:

    • stereoselective - can differentiate between enantiomers and between prochiral substrates
    • regioselective - can differentiate between identical functional groups in a single substrate
    • chemoselective - can differentiate between different functional groups in a substrate (such as between a hydroxyl group and an amine for an acylation reaction)

    Enzyme in anhydrous organic solvents are useful (from a synthetic point) not only since new types of reactions can be catalyzed (such as transesterification, ammonolysis, thiolysis) but also because the stereoselectivity, regioselectivity, and chemoselectivity of the enzyme often changes from activities of the enzyme in water.

    Organic reactions are usually conducted in organic solvents, since many organic molecules react with water, and the reagents and products are usually not soluble in water. In a manner analogous to using an enzyme as a heterogeneous catalyst in nonpolar solvent, Sharpless is pioneering a technique to conduct organic reactions in water. They (Narayan et al.) have shown that many unimolecular and bimolecular reactions occur faster in water than in organic solvents. As in enzyme catalysis in nonpolar solvent, the reactions must be mixed vigorously to disperse reactants in micro-drops (a suspension) in water, greatly increasing the surface area that might allow water to act on transition states or intermediates to stabilize them through hydrogen bonding. They called these reactions "on water" reactions since reactants usually float on water. They have performed cycloadditions, alkene reactions, Claisen rearrangments, and nucleophilic substitution reactions using this process. One cycloaddition reaction went to completion in ten minutes at room temperature, compared to 18 hours in methanol and 120 in toluene. Adding nonpolar solvent at certain times greatly increased the rate of the reaction.

    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, as shown in Figure (\PageIndex{24}\) 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
    Figure (\PageIndex{24}\): DNA of the CRISPR operon

    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 eukaryotes. The hope is that CRISPR technology will give us a precise and incredibly cheap way to do gene therapy in diseased cells and organisms. Given its role in transforming our ability to edit the genome and potentially cure genetically-based diseases, we will offer a detailed explanation of its mechanism.

    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.

    Apo- and Holo-Cas 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 Figure (\PageIndex{25}\) 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).

    Figure (\PageIndex{25}\): Apoenyzme Cas9 (without bound RNA and DNA (4cmp)

    A close up view showing the two catalytic sites is shown in Figure (\PageIndex{26}\) below.

    Figure (\PageIndex{26}\): Two catalytic sites in Cas9

    Figure \(\PageIndex{27}\) below shows an interactive iCn3D model of Streptococcus pyogenes Cas9 in complex with guide RNA and target DNA (4OO8) (long load time). The Cas9 enzyme is shown as a gray transparent surface with an underlying cartoon rendering. The DNA is shown as colored sticks. The RNA is shown as a cyan cartoon.n.

    Streptococcus pyogenes Cas9 in complex with guide RNA and target DNA 4OO8.png

    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{27}\): Streptococcus pyogenes Cas9 in complex with guide RNA and target DNA (4OO8). (Copyright; author via source).
    Click the image for a popup or use this external link: (long load time)

    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 Figure (\PageIndex{28}\) below.

    Figure (\PageIndex{28}\): Structure of the holoenzyme Cas9 (ternary complex)with bound guide RNA and DNA

    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 Figure (\PageIndex{29}\) below. The importance of this change will be described later.

    combo CRISPR Surf Conf Change
    Figure (\PageIndex{29}\): Distance between D435 and E 944/945 going from the apo-Cas9 (left) to the holo-Cas 9 (right) enzyme

    Figure (\PageIndex{30}\) 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.

    Figure (\PageIndex{30}\): 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

    Figure (\PageIndex{31}\) below shows an expanded view of the ternary complex.

    Figure (\PageIndex{31}\): Expanded view of the ternary complex of Cas9 with guide RNA and DNA

    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. Figure (\PageIndex{32}\) below shows the Args:PAM interaction (pdb code 4un3)

    Figure (\PageIndex{32}\): Args:PAM interaction in holo-Cas9 (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".

    Figure (\PageIndex{33}\) below show an animation that illustrates 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
    Figure (\PageIndex{33}\): Animation illustrating conformational changes going from the apo Cas 9 to the binary Cas 9:sgRNA complex to the ternary Cas 9: sgRNA: target DNA complex

    A potential abbreviated catalytic mechanism for the Ruv nuclease domain is shown in Figure (\PageIndex{34}\) 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.

    Figure (\PageIndex{33}\): Abbreviated catalytic mechanism for the Ruv nuclease domain of Cas9

    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.

    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 as shown in panel A in Figure (\PageIndex{33}\). 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 Panel B in Figure (\PageIndex{33}\). 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 as shown in Panel C of Figure (\PageIndex{33}\).

    Figure (\PageIndex{33}\): 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 as shown in panel A of Figure (\PageIndex{33}\) that is induced into the 'closed' and biologically active conformation upon substrate binding.

    Figure (\PageIndex{33}\): 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):e1003521and Figure C from: Snodgrah

    Within the AK protein structure, there is a core domain and two smaller domains called the LID and NMP as shown in Figure (\PageIndex{34}\) below. 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 (\PageIndex{34}\): 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

    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 (\PageIndex{35}\): 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. Exampoles of staggers and blunt end cuts by restrictions enzymes are shown in Table \(\PageIndex{8}\) below.

    EcoR1 EcoRI_restriction_enzyme_recognition_site.svg
    Sma1 SmaI_restriction_enzyme_recognition_site.svg

    Table \(\PageIndex{8}\): Staggered and blunt end cut sequences by EcoR1 and Sma1

    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 as shown in Figure (\PageIndex{36}\):. 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 (\PageIndex{36}\): 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 chiral center 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 (\PageIndex{37}\) below.

    Figure (\PageIndex{37}\): A General Mechanism for DNA Cleavage by EcoRI and EcoRV. An activated water molecule attacks the phosphorous in-line with 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 as shown in Panel A of Figure (\PageIndex{38}\) below (from: Pingoud, A., Wilson, G.G., and Wende, W. (2014) Nuc Acids Res 42(12):7489-7527.. 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 (\PageIndex{38}\): 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

    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. As we saw previously, ssRNA can adopt seconary, tertiary and quaternary structures with protein. The fact that they could act as catalyst, as enzymes, hence should not be surprising. We will discuss the mechanism of ribozymes in its own section (X.X)

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    6.5: Enzymatic Reaction Mechanisms is shared under a not declared license and was authored, remixed, and/or curated by Henry Jakubowski.

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