Kinetic Analysis of Class II Lanthipeptide Maturation Catalyzed by ProcM

By

Yunhong Wang

Thesis for the

Degree of Bachelor of Science in Biochemistry

College of Liberal Arts and Sciences University of Illinois

Urbana-Champaign, Illinois

2015

  i  

Abstract:

Lanthipeptides are a class of ribosomally synthesized and post-translationally modified

peptide natural products (RiPPs) that are characterized by intramolecular thioether ring structures.

These cross-links endow lanthipeptides with various biological activities, including antibiotic

activity against multiple bacterial pathogens. In the hopes of manipulating their antibiotic

activity and engineering them into novel antibiotics, it is critical to understand how these ring

structures are enzymatically constructed. A special class II lanthipeptide synthtase, ProcM, has

received much attention recently because of its extremely high substrate tolerance. This enzyme

installs thioether linkages with drastically different topologies into a total of 29 different natural

substrates. In this work, the maturation process of one of these 29 substrates (ProcA2.8) was

characterized in detail by performing LC/ESI-MS kinetic studies on a set of ProcA2.8 variant

peptides. From the numerical simulation of the experimental data, kinetic parameters for

individual ProcM-catalyzed post-translational modifications were determined. This analysis

provided much insight into how the maturing structure of the ProcA2.8 precursor peptide

modulates the kinetics of the overall maturation process. On the basis of this data, I suggest a

model for ProcM function where the kinetic parameters for the different modifications are

governed largely by the molecular binding interactions between the ProcA2.8 core peptide and

the active sites of the ProcM enzyme.

  ii  

Acknowledgement:

First and foremost, I would like to thank Professor Wilfred van der Donk with great

appreciation for the opportunity of working in his lab on lanthipeptides research since Fall 2012.

Credits also go to my mentor, Christopher Thibodeaux who has provided extensive guidance

throughout my three-year research as well as academic life. In addition, he has designed such an

innovative and interesting project. Without his help, my thesis work is not possible. Also, I

would like to thank other members in van der Donk lab for creating an optimistic working

environment. It has been a joy working with them.

Besides, I could not achieve my accomplishments in research as well as academic life

without my comrades who always fight together with me days and nights. I hereby thank

Bingyan Wu, Yanshu Guo, Fei Wang, Ziqiao Ding, Kevin Gill and Daniel Lim. Solving

problems and overcoming difficulties with them have become unforgettable memories. In

addition, I would like to thank my other friends who accompanied me in this four-year journey. I

hereby thank Hanqing Ye, Xizhu Wang, Lunan Sun, Liwei Song, Junji Zhu, Yijian Gu, Ziyan

Zhou, Yuchen Chai, Feng Shan and Weikun Xiao for sharing happiness and sadness with me.

They have taught me important lessons in my life and made Urbana-Champaign like home. Our

memories are engraved in my mind.

Last but not least, I would like to thank my parents, Yuefei Wang and Lingyun Sheng,

who made everything possible and always supported me financially and emotionally.

  iii  

Collaboration:

This thesis work was accomplished under direct instruction from Dr. Christopher

Thibodeaux, who provided extensive guidance throughout the project.

  iv  

Table of Contents:

Abstract ............................................................................................................................................ i Acknowledgement and collaboration ............................................................................................. ii Table of contents ............................................................................................................................ iv

Introduction ..................................................................................................................................... 1 Materials and Methods .................................................................................................................. 12 Results and Discussion ................................................................................................................. 20 Conclusion .................................................................................................................................... 47 Bibliography ................................................................................................................................. 49

  1  

Introduction:

Overview of Lanthipeptides

Since the discovery of the first antimicrobial compound, penicillin, by Alexander

Fleming in 1928, thousands of antibiotics have been discovered or synthesized. Antibiotics

revolutionized medicine in the 20th century due to their success in the treatment of bacterial

infections and in chemotherapy. However, with the increasing use (and misuse) of antibiotics,

many bacterial strains have mutated to develop resistance against multiple antibiotics, with some

strains even developing resistance against all classes of commercially available antibiotics. To

prevent future outbreaks of epidemic diseases, novel antibiotics are urgently needed to replenish

the dwindling supply of effective antibacterial agents. Among the many classes of microbial

secondary metabolites, lanthipeptides are a promising class of peptide natural products that

exhibit antimicrobial activity and avoid common bacterial resistance mechanisms.

The most well known industrial application of lanthipeptides is the use of nisin in the

food industry (Figure 1). Nisin is active against a range of food-borne pathogens, including

Bacillus and Listeria species (30), and has been widely used as a food preservative in many

products such as processed cheese and meat for over fifty years. Remarkably, despite its

widespread use for many years, stable bacterial resistance to nisin has not been experimentally

documented (2). The promising antimicrobial properties of lanthipeptides such as nisin, suggest

that this class of compounds could become a vital component of our future antimicrobial arsenal.

As such, a thorough understanding of the biochemical properties of lanthipeptide biosynthetic

enzymes could provide valuable information for the bioproduction and/or engineering of novel

lanthipeptide structures that could potentially help to combat antibiotic resistance.

  2  

Lanthipeptides are

ribosomally synthesized

and post-translationally

modified peptides (RiPP)

natural products that often

exhibit antibiotic activity.

Lanthipeptides can exhibit

multiple modes of action,

typically involving the

inhibition of bacterial cell wall biosynthesis and/or the formation of pores in the bacterial plasma

membrane (3-6). Specifically, lanthipeptides often bind to lipid II (an important precursor of the

bacterial cell wall) and prevent its utilization by the transpeptidase and transglycosylase enzymes

of cell wall biosynthesis. This interferes with the formation of the cross-linked peptidoglycan

network that is critical for the structural integrity of the bacterial cell wall (3). Some

lanthipeptides (such as nisin) also form pores in the bacterial cell membrane, leading to a loss of

membrane potential and the rapid death of the target cell. Thus, the biological targets of

lanthipeptides are essential for bacterial survival and are highly conserved through evolution, and

it is thought that these features contribute to the effectiveness of lanthipeptides at evading

common bacterial resistance mechanisms. However, despite the impressive antimicrobial

characteristics of many lanthipeptides, their clinical usage is often limited by their poor

pharmacokinetic properties. Therefore, there is a great need to discover and characterize new

lanthipeptides, and to engineer existing compounds into more promising drug leads.

Figure 1. Chemical structure of nisin A. The mature core peptide of nisin contains one lanthionine ring (red), four methyllanthionine rings (blue), a Dha residue (red) and a Dhb residue (purple). These characteristic structures contribute to antimicrobial activity (17).  

  3  

Biosynthesis of Lanthipeptides

Like all other RiPPs, lanthipeptides are genetically encoded and are first expressed as an

inactive linear precursor peptide, generically termed LanA. LanA precursors contain an N-

terminal leader sequence (called the leader peptide) as well as a C-terminal sequence that harbors

the posttranslational modifications (termed the core peptide). Among other functions, the leader

peptide serves as a critical binding element for the lanthipeptide synthetases that catalyze the

modifications of the core peptide (7). Lanthipeptide synthetases are responsible for the

biosynthesis of the lanthionine (Lan) and methyllanthionine (MeLan) moieties that typify

lanthipeptide structure (Figure 2). The chemical mechanism for the formation of these thioether

rings involves dehydration of serine (Ser) and threonine (Thr) residues in the core peptide to

form dehydroalanine (Dha) and dehydrobutyrine (Dhb) residues, respectively, followed by the

intramolecular Michael-type addition of cysteine (Cys) residues onto a subset of Dha and Dhb

residues to form Lan and Melan thioether linkages, respectively (Figure 2). In some

lanthipeptides, the Dha and Dhb residues are not subjected to Michael addition by Cys residues.

  4  

Following modification of the core peptide, the mature precursor peptide is exported

from the producing cell and the leader peptide is proteolytically removed to unmask the

biological activity of the lanthipeptide. In all cases that have been studied, the thioether rings are

critical for the biological activity of lanthipeptides, providing both target binding specificity and

stability towards degradation by target cell proteases (4, 5). Therefore, in order to manipulate

lanthipeptide synthetases for the engineering of novel antibiotics, it is important to study the

formation of these ring structures in detail.

Lanthipeptide Synthetases

Based on differences in the requisite biosynthetic machinery (Figure 3), lanthipeptides

can be categorized into four distinct classes (8-11). In class I lanthipeptides, such as nisin, the

dehydratase and cyclase activities needed for thioether biosynthesis are encoded on two separate

polypeptides. The dehydration step is catalyzed by the LanB enzyme, and the cyclization step is

catalyzed by the LanC enzyme (12). For the class II, III, and IV lanthipeptides, a single

Figure 2. Chemical mechanism of class I and class II lanthipeptide post-translational modification. For class I lanthipeptides, LanB carries out dehydration via glutamylation. LanC is responsible for cyclization. For class II lanthipeptides, bi-functional LanM carries out dehydration via phosphorylation as well as cyclization.

  5  

multifunctional enzyme

catalyzes all of the steps

needed for thioether

biosynthesis (8, 13). For

class II lanthipeptides, the

posttranslational

modifications are installed

by a LanM enzyme that contains an N-terminal dehydratase domain and a C-terminal cyclase

domain. In contrast, class III and class IV lanthipeptides are synthesized by the trifuntional

enzymes LanKC (14, 15) and LanL (9), respectively, both of which contain an N-terminal lyase

domain, a central kinase domain and a C-terminal cyclase domain.

Although all four lanthipeptide biosynthetic systems have the ability to install thioether

rings in their substrates, the domain function and organization of all four enzyme classes differ.

Other than the ligands to the catalytic Zn atom, LanC and the cyclase domains of the class II-IV

lanthipeptide synthetases share very little (but still detectable) amino acid sequence homology.

Interestingly, the cyclase domain of LanKC (the class III synthetase) lacks the otherwise

conserved Zn ligands. Phylogenetic analysis of the LanC cyclase and cyclase domains of class

II-IV synthetases suggest that LanM and LanL have evolved independently from LanC (11).

Moreover, the enzymes that are responsible for the dehydration reaction in each of the four

lanthipeptide synthetases are even more evolutionarily divergent. For class I lanthipeptides, the

dehydration is carried out by a separate enzyme (LanB), which has recently been shown to

catalyze the dehydration of Ser/Thr residues via a glutamylated intermediate using glutamylated-

tRNAGlu borrowed from primary metabolism (Figure 2) (16, 17). In contrast, the class II, III, and

Figure 3. Functional domains in lanthipeptide synthetases. The conserved zinc-binding residues are highlighted with purple lines in the cyclase domains. (11)  

  6  

IV synthetases catalyze dehydration in an ATP-dependent manner, wherein the γ-phosphate of

ATP is first transferred to the Ser/Thr residue to be dehydrated to generate a phosphopeptide

intermediate (Figure 2). Phosphorylation (and glutamylation in the case of class I lanthipeptides)

provides a good leaving group for the subsequent base-catalyzed elimination step to complete the

dehydration. The N-terminal dehydratase domain of LanM shares no amino acid sequence

similarity with the class I LanB dehydratase (or with any other protein of known function),

perhaps suggesting a novel structural fold that has evolved specifically for catalyzing

lanthipeptide dehydration. In contrast to the class I and class II systems, the LanKC and LanL

enzymes carry out the dehydration reaction using kinase and lyase catalytic domains that share

homology with proteins of known function. The different classes of lanthipeptide synthases

provide a striking example of convergent evolution, where different structural folds have been

repurposed and adapted over evolutionary time in order to synthesize the same chemical

structure (i.e. (methyl)lanthionine rings). This convergent evolution underscores the importance

of the lanthipeptide moiety in biological systems (11).

Biochemical Studies of LanM Enzymes

Mechanistically, the LanM enzymes (the subject of this thesis) are the best-studied

lanthipeptide synthetases. Previous biochemical studies on the lacticin 481 biosynthetic system

performed by our group have revealed conserved amino acid residues that play roles in the

dehydration and cyclization activities of LanM enzymes. You et al. identified amino acid

residues that are important in the dehydration carried out by LctM on its substrate, LctA (18).

They first conducted sequence alignments on nine members of class II lanthipeptide synthetases,

and identified the fully conserved amino acid residues among them. They then mutated these

  7  

conserved residues and incubated the mutant enzymes with wild type LctA. From analysis of the

reaction products, they then deduced functions for the conserved residues. The results of this

study are presented in Figure 4.

In the phosphorylation step, either Asp242 or Asp259 is believed to serve as a base to

deprotonate the Ser/Thr hydroxyl for attack on the γ-phosphate. When either of these residues is

mutated, no modification of the LctA precursor peptide was observed. These two residues may

interact through a hydrogen bond that could serve to activate one of the two residues as a base to

deprotonate the Ser/Thr hydroxyl group. Similarly, Lys159 also appears to play a role in the

Ser/Thr phosphorylation step, perhaps by stabilizing the developing negative charge on the ADP

leaving group during phosphoryl transfer to Ser/Thr. Lys159 may also stabilize negative charge

Figure 4. Putative chemical mechanism of dehydration (A) and cyclization (B) by LctM. In the dehydratase active site, Asp242, Asp259 and Arg399 play important roles in the phosphorylation and elimination steps. In the cyclase active site, Cys781, Cys836 and His837 coordinate the catalytic Zn2+ while His725 likely protonates Cα (18, 20, 31).

  8  

formation on the phosphate leaving group during the elimination step. Finally, Arg399 and

Asp259 both seem to play roles in the phosphate elimination step. The R399M mutant

accumulates phosphopeptide intermediates, suggesting that the mutant is deficient in phosphate

elimination. As mentioned above, the Asp242 and Asp259 mutants were devoid of catalytic

activity; however, when the D242N mutant was presented with a phosphorylated peptide

substrate, the phosphate was eliminated to produce the dehydrated residue. In contrast, the

D159N mutant was unable to catalyze phosphate elimination with this substrate. Cumulatively,

these studies implicate Asp259 as the base that deprotonates the Cα atom of the pSer/pThr

residue during the phosphate elimination step. In this model, Arg399 (which likely carries a

positive charge due the high pKa of ~ 12 of the guanidinium side chain) may form an ion pair

with Asp259, lowering the pKa of the latter to facilitate its function as the catalytic base (18).

Previous studies have also shown that lanthipeptide cyclases function in a zinc-dependent

manner (19). Paul et al. has shown that in LctM, the three amino acid residues, Cys781, Cys836

and His837, coordinate the zinc ion that plays a role in the cyclization event. Through

mutagenesis studies, they have shown that the presence of Cys781 and Cys836 are required for

correct enzymatic cyclization of LctA but not the dehydration of LctA, which suggests that

dehydration and cyclization are independent events catalyzed by different domains of the same

enzyme (20). The zinc atom in the active site likely serves to deprotonate and position the Cys

thiols for nucleophilic attack on the dehydroamino acids (Figure 4). A conserved His residue

(His725 in LctM) could serve as the active site acid that protonates the incipient enolate to

complete the transformation. Structural analysis of fully modified lanthipeptides has shown that

lanthipeptide cyclase domains typically install methyllanthionine residues with the (2S, 3S, 6R)

configuration and lanthionine residues with the (2S, 6R) configuration (Figure 2) (4, 21),

  9  

suggesting that attack of the Cys thiol and protonation of the Dha/Dhb double bond occur from

opposite faces of the planar alkene moiety of the Dha/Dhb group. Although the precise

mechanisms that govern the regioselectivity of thioether ring formation by lanthipeptide

synthetases are currently unknown, it is believed that the ring topology present in the final

compound is governed by some combination of the primary amino acid sequence of the core

peptide and/or the specific molecular interactions between the peptide and the synthetase.

ProcM is an unusual LanM enzyme

Recently, a very unusual class II lanthipeptide biosynthetic system was identified in the

marine cyanobacterium, Prochlorococcus marinus MIT9313 (22). In this organism, a single

LanM enzyme encoded in the genome (ProcM) carries out the post-translational modification of

30 different LanA peptide precursors (11, 22), providing an unusual example of natural

combinatorial biosynthesis. In contrast to most LanM enzymes, which process a single substrate

into a single product with a defined thioether ring topology, ProcM installs many different ring

topologies into core peptides with hypervariable sequences (8, 22). ProcM shares the conserved

residues found in LctM and other class II lanthipeptide synthetases; however, the zinc in the

cyclase domain of ProcM has an unusual coordination sphere composed of three Cys ligands,

instead of the typical Cys2His coordination geometry found in most class II synthetases. This

unusual zinc coordination could be related to the ability of ProcM to install multiple thioether

topologies (11, 32).

The reaction of ProcM with ProcA2.8 is one of the best-studied natural ProcM reactions.

In the ProcM/ProcA2.8 reaction (Figure 5), ProcM catalyzes dehydration of Ser9 and Ser13 in

the ProcA2.8 core peptide, followed by the formation of two non-overlapping lanthionine rings

  10  

(formed by attack of Cys3

on Dha9 and by attack of

Cys19 on Dha13). Recent

mechanistic studies on the

ProcM/ProcA2.8 reaction

demonstrated that both the

dehydrations and the

cyclizations preferentially occur in a C- to N-terminal direction (i.e. Ser13 is dehydrated before

Ser9, and ring B is formed before ring A) (24). Kinetic studies on these modifications have

shown that both dehydrations tend to be completed prior to the onset of cyclization, and that the

cyclization of ring A is the slowest overall step in the pathway (25). Furthermore, the two

cyclization events occur at markedly different rates, with ring B formation being approximately

10 fold faster than ring A formation (25). Preliminary kinetic studies on other ProcM substrates

have shown that the first ring is typically installed quickly, while the remaining rings are

installed more slowly. This observation is consistent with a model where ProcM has not evolved

substrate specificity or kinetic efficiency for any one of its 30 substrates. The kinetic properties

of ProcM contrast markedly with the kinetic properties of the substrate selective class II

synthetase HalM2, which was found to very efficiently and quickly catalyze the maturation of its

natural substrate, HalA2 (25). The poor cyclization kinetics of ProcM are perhaps expected for

an enzyme that is charged with installing so many different thioether ring topologies.

Despite the insights these previous studies provided into the catalytic properties of ProcM,

the mechanism by which ProcM maintains its relaxed substrate specificity is largely unknown.

The current hypothesis is that a series of conformational changes and protein-protein

Figure 5. Schematic representation of ProcM catalyzed ProcA2.8 maturation (25)  

  11  

interactions between LanM enzymes and their LanA substrates govern the selection of

posttranslational modification (PTM) sites, as well as the timing of the various PTMs. In this

model, the changing structure of the LanA substrate as it matures into the final product could

potentially influence the kinetic parameters and, hence, the preferred order of subsequent steps in

the biosynthetic pathway. In order to test this model, I have constructed a series of ProcA2.8

variant peptides designed to allow us to measure the kinetic parameters of each ProcM-installed

modification in the presence and absence of other PTMs. In this manner, the effects of ProcM-

installed PTMs on the kinetics of subsequent modifications can be directly probed. The studies in

this thesis are expected to significantly advance our understanding of the factors that govern

LanM substrate specificity and catalytic efficiency, which, in turn, may facilitate the use of these

impressive enzymes for the engineering of novel lantibiotics with new and/or improved

biological activities.

  12  

Materials and Methods:

Preparation of ProcA2.8 mutant peptides

Cloning:

The Escherichia coli expression constructs encoding the ProcA2.8 variant peptides were

constructed by GeneScript using the procA2.8/pET15b vector as the template (22). Each mutant

peptide gene (Table 1) was inserted into the NdeI/XhoI restriction sites of pET15b to generate a

construct encoding an N-terminal His6-affinity tag. In each of the ProcA2.8 mutants, one or more

of the amino acid residues that are involved in the post-translational modifications (Cys3, Ser9,

Ser13 and Cys19) were mutated into alanine residues. The corresponding codons for the Cys and

Ser residues were mutated into the most frequently utilized alanine codon in E. coli (GCG). The

preparation processes for all of the ProcA2.8 mutants were the same unless otherwise stated.

Each of the procA2.8/pET15b constructs was dissolved in 50 µL of H2O. Escherichia coli BL21

(DE3) chemically competent cells (50 µL) were transformed with 1 µL of the resulting

recombinant vector using heat shock for 30 s in a 42 °C water bath immediately followed by a 2

min incubation on ice. The cells were inoculated in 1 mL of Luria broth (LB) medium in a 37 °C

shaker for 1 h. The cell culture was then centrifuged at 13,000×g for 2 min. A sample of 900 µL

of the supernatant was removed and discarded. The cell pellet was resuspended in the remaining

100 µL of media, and all of the cells were plated on pre-warmed LB-ampicillin (100 µg/mL)

plates. The plates were incubated at 37 °C overnight.

  13  

Table 1. Nucleotide sequence of procA2.8 variant genes

Variant Nucleotide sequence of core peptide (5´-3´) ProcA2.8-wt GCGGCCTGTCATAACCATGCTCCATCTATGCCTCCATCCTATTGGGAGGGTGAGTGCTAA

ProcA2.8-S1 GCGGCCGCGCATAACCATGCTCCATCTATGCCTCCAGCGTATTGGGAGGGTGAGGCGTAA

ProcA2.8-S2 GCGGCCGCGCATAACCATGCTCCAGCGATGCCTCCATCCTATTGGGAGGGTGAGGCGTAA

ProcA2.8-S1S2 GCGGCCGCGCATAACCATGCTCCATCTATGCCTCCATCCTATTGGGAGGGTGAGGCGTAA

ProcA2.8-C1S1 GCGGCCTGTCATAACCATGCTCCATCTATGCCTCCAGCGTATTGGGAGGGTGAGGCGTAA

ProcA2.8-S2C2 GCGGCCGCGCATAACCATGCTCCAGCGATGCCTCCATCCTATTGGGAGGGTGAGTGCTAA

ProcA2.8-C1S1S2 GCGGCCTGTCATAACCATGCTCCATCTATGCCTCCATCCTATTGGGAGGGTGAGGCGTAA

ProcA2.8-S1S2C2 GCGGCCGCGCATAACCATGCTCCATCTATGCCTCCATCCTATTGGGAGGGTGAGTGCTAA

The nucleotide sequence for the leader peptide is not shown. Overexpression:

Single colonies of transformants were selected from the LB-Amp plate and were grown

in 20 mL of LB medium supplemented with 100 µg/mL ampicillin in a 37 °C shaker overnight.

The pre-culture was then centrifuged at 4,000×g for 10 min at 4 °C. The spent LB medium was

discarded and the cell pellet was carefully resuspended in fresh LB medium. This step aimed to

remove the secreted β-lactamase, which degrades ampicillin and decreases the selection pressure

on E. coli transformants that carry the pET15b vector. The resuspended cells (20 mL) were

added to 2 L of LB medium supplemented with 100 µg/mL ampicillin, and the culture was

incubated in a 37 °C shaker until the optical density at 600 nm (OD600) reached approximately

0.5, at which point isopropyl-β-D-1-thiogalactopyranoside (IPTG) was added to a final

concentration of 250 µM to induce the cell growth. The cell culture was then grown at 18 °C

overnight. Cells were harvested by centrifugation at 8,000×g for 15 min at 4 °C. The cell paste

was stored at -80°C until use.

  14  

Peptide Purification:

The cell paste was resuspended in ~20 mL of LanA Lysis/Wash Buffer (4 M guanidine

hydrochloride, 20 mM NaH2PO4, 500 mM NaCl, 30 mM imidazole, 20% glycerol, pH 7.5 ) and

lysed by sonication (35% amplitude, 4.9 s pulse on, 9.9 s pulse off, 15 min). The samples were

centrifuged at 14,000×g for 30 min at 4 °C. Immobilized metal affinity chromatography (IMAC)

was carried out to purify the peptide using a HiTrap chelating HP nickel affinity column (GE

Helthcare). The HiTrap resin was charged with 10 mL of 0.1 M NiSO4, washed with 10 mL of

H2O and equilibrated with 30 mL of LanA lysis/wash buffer. The supernatant from the

centrifuged crude cell lysate was loaded onto the column. The column was washed with 50 mL

of LanA lysis/wash buffer to remove non-specifically bound protein contaminants. The desired

ProcA2.8 mutant peptides were eluted with LanA elution buffer (4 M guanidine hydrochloride,

20 mM Tris, 100 mM NaCl, 250 mM imidazole, pH 7.5).

The imidazole was removed by running the IMAC elute sample through a Vydac 1 mL

C4 solid phase extraction (C4-SPE) column. A 1 mL portion of the IMAC elute was loaded onto

the column. The column was washed with 3 mL of 0.1% trifluoroacetic acid (TFA) in H2O, and

eluted with 2 mL of 0.1 TFA in 80% acetonitrile (MeCN). The C4-SPE elutes were pooled, flash

frozen in liquid nitrogen, lyophilized, redissolved in H2O water and stored at -20 °C. To remove

the His6-affinity tag, the crude peptides were treated with 50 units of high purity grade bovine

thrombin in 25 mM potassium phosphate buffer (pH 7.0) at 4 °C for 24 h. The thrombin-

digested ProcA2.8 mutant peptide preparations were further purified by reversed phase high

performance liquid chromatography (HPLC) on a Shimadzu Prominence Preparative Liquid

Chromatography instrument equipped with a Phenomenex Luna C5 semiprep column (250×10

mm, 10µ, 100 Å) at a flow rate of 7 mL/min and a gradient of 2-62% solvent B over 30 min and

  15  

62-100% solvent B over 5 min (solvent A = 0.1% TFA in H2O; solvent B = 80% MeCN in H2O,

0.1% TFA). Finally, peptides were flash frozen in liquid nitrogen, lyophilized, redissolved in

H2O water and stored at -20° C until use.

Preparation of ProcM synthetase:

Cloning and Overexpression:

The procM gene was cloned into the Nde1/Xho1 sites of pET28 as described previously

(22). E. coli BL21 (DE3) cells were transformed with the recombinant vector by heat shock (30 s

in 42 °C water bath, 2 min on ice). The ProcM protein was expressed following the same

procedure that was used for the overexpression of ProcA2.8 mutants, except that the antibiotic

used was 30 µg/mL kanamycin. The cell paste was harvested by centrifugation at 8,000 ×g for

15 min at 4 °C and was stored at -80 °C.

Protein Purification:

The cells were dissolved in lysis/wash buffer (25 mM Tris, 500 mM NaCl, 20 mM

imidazole, 10% glycerol, pH 8.0), lysed by sonication (35% amplitude, 4.9 s pulse on, 9.9 s pulse

off, 10 min), and centrifuged at 14,000×g for 30 min at 4 °C. The supernatant was loaded onto an

equilibrated 5 mL Hi-Trap Ni-NTA column, washed with 50 mL of lysis/wash buffer and eluted

with 15 mL of elution buffer (25 mM Tris, 500 mM NaCl, 200 mM imidazole, 10% glycerol, pH

8.0). The 15 mL elute was concentrated to ~ 3 mL using an Amicon Ultra Centrifugal Filter

device with a 50 kDa MW cutoff (Milipore). The 3 mL sample was further purified by gel

filtration chromatography (Superdex 200, 1.5 × 60 cm, GE Healthcare) at 4 °C, using a 1

mL/min flow rate and an isocratic elution (25 mM Tris, 500 mM NaCl, 10% glycerol, pH 8.0).

  16  

The ProcM protein typically elutes in three peaks (aggregate, then oligomer, followed by

monomer). Only the monomer fraction was collected and further concentrated using a second

Amicon centrifugal filtration device (50 kDa). Aliquots for one time use were flash frozen in

liquid nitrogen, and stored at -80 °C.

Kinetic Assays:

The kinetic experiment for each of the ProcA2.8 mutants with ProcM was performed in

duplicate. Before carrying out the ProcM/ProcA2.8 mutant reactions, 2 µM of His6-tagged

ProcM synthetase and 80 µM of ProcA2.8 mutants were separately pre-incubated for 1 h at

25 °C in 750 µL of reaction mixture containing 5 mM ATP, 5 mM MgCl2, 100 mM HEPES (pH

7.5) and 0.1 mM TCEP. Pre-incubation allows the enzyme and substrate to fully equilibrate with

the reaction environment prior to initiating the reaction. To initiate the ProcM/ProcA2.8 reaction,

the two portions of the mixture were thoroughly mixed together, resulting in a reaction mixture

containing 1 µM His6-tagged ProcM, 40 µM of ProcA2.8 peptide, and the other reaction

components listed above. An 80 µL aliquot was removed from the reaction at 17 desired time

points (2 min, 4 min, 6 min, 8 min, 10 min, 12 min, 15 min, 18 min, 20 min, 30 min, 45 min, 60

min, 75 min, 90 min, 120 min, 180 min, and 240 min) and was quenched into 900 µL of quench

buffer (111 mM citrate pH 3.0, 1.11 mM EDTA). The quenched reaction aliquots were stored on

ice until the last time point was quenched.

Derivitization of Kinetic Assay Time Points:

After the last time point was quenched, to keep the thiols in their reduced state, all of the

reaction aliquots were spiked with 100 µL of 100 mM TCEP and incubated for 10 min at 25 °C.

  17  

At this point, the pH of each quenched time points was below 3.5. The pH of each aliquot was

then adjusted to approximately pH 6.3 by adding 35-40 µL of 5 M NaOH. To alkylate free Cys

thiols, 11 µL of 1 M N-ethylmaleimide (NEM) in ethanol was added into each quenched reaction

aliquot and incubated at 37 °C for 10 min. After NEM alkylation, the aliquots were quenched

with 11 µL of 100% TFA and were purified by C4-SPE column. The C4-SPE column was first

equilibrated with 3 mL of 0.1% TFA in H2O. The peptide sample was loaded onto the column,

washed with 3 mL of 0.1% TFA, and eluted with 2 mL of 0.1% TFA in 80% MeCN. The eluates

were flash frozen in liquid nitrogen, lyophilized, and stored as powder at -20 °C until LC-ESI-

MS analysis.

Liquid Chromatography Electrospray Ionization Mass Spectrometry (LC-ESI-MS):

For LC-ESI-MS analysis, lyophilized peptides were dissolved in 50 µL of H2O. The

concentrations of the peptides were then estimated by their absorption at 280 nm (ε280= 7115 M-

1cm-1). Peptide samples were then diluted to final concentrations of 10 µM in LC-ESI-MS

injection solvent (50% MeCN in H2O, 0.1% formic acid) and were analyzed within 12 h at 8 °C

to prevent solvent evaporation. Peptide samples (5 µL) were injected into an Acquity UPLC

BEH C8 1.7 µM column (1 x 100 mm) attached to a quadrupole/time-of-flight (Q/TOF) Synapt

mass spectrometer (Waters). To minimize systematic errors, all samples were injected in a

randomized order and 10 µL H2O blanks were run between each sample to ensure that all peptide

from the previous sample had eluted from the column. Before injection, the instrument was

calibrated using a 0.1% phosphoric acid standard. The peptide was eluted using the following

gradient: 2 – 100% solvent B over 20 min at a flow rate of 0.090 mL/min (solvent A = 0.1%

formic acid in H2O, solvent B = 100% MeCN, 0.1% formic acid). Under these conditions, all

  18  

peptides analyzed in this study eluted as a broad peak centered at about 7 min. The Synapt

instrument settings were as follows: positive ion mode, capillary voltage = 3.0 kV, cone gas = 20

L/h, desolvation gas = 500 L/h, source temperature = 110 °C, desolvation temperature = 150 °C.

Data were collected in centroid mode with an extended linear range without precursor ion

selection. The TOF detector was set to an m/z window of 800 – 1800 Da, and a 1 s scan time was

used for data acquisition. The m/z values for the ions of interest were determined by summing

the mass spectra at full width at half maximum (FWHM) of the main chromatographic peak

observed in the total ion chromatogram (TIC) of each LC injection. Extracted ion

chromatograms (EICs) were then generated by applying a mass window of variable size (1.5 –

2.5 Da, depending on the charge state and the intensity of the ion signal) around the center of the

most intense isotope peak for each ion of interest. The EICs were integrated using MassLynx

software (Waters) to generate EIC peak areas. A correction of 50 peak area units was subtracted

from each EIC peak area in order to account for background. The EIC peak areas were then used

for calculation of the fractional abundances of the peptides of interest as described below.

Semiquantitative Analysis of Mass Spectrometry Data:

The extracted ion chromatograms (EICs) were used to calculate the fractional abundance

of each ProcA2.8 species of interest using a procedure analogous to that reported by Kelleher

and co-workers (27, 28). Using Equation 1, the fractional abundance of peptide species X in

charge state i (fX,i) was determined as the ratio of the EIC peak area for species X in charge state i

(AX,i) to the sum of the EIC peak areas for all the relevant peptide species in charge state i (Ai).

𝑓!,! =!!,!!!

Equation 1

  19  

The weight of charge state i (wi) is calculated by Equation 2 as the ratio of the sum of the EIC

peak areas for all relevant peptides in charge state i (Ai) to the sum of the EIC peak areas for all

of the relevant peptide species in all of the charge states observed in the mass spectrum (Atotal)

𝑤!!

!!!!"!#$

Equation 2

Finally, the charge state-weighted fractional abundance of peptide species was calculated using

Equation 3, where the sum is over all of the charge states (i) observed in the mass spectrum.

𝐹! = 𝑤!𝑓!,! Equation 3

Time dependent changes in the fractional abundances of the relevant reaction species were then

numerically simulated by my advisor (Dr. Christopher Thibodeaux) using Kintek Explorer (25,

26) as described previously (23). This process enabled us to evaluate kinetic models for

ProcM/ProcA2.8 biosynthesis and to determine rate constants for the ProcM-catalyzed

posttranslational modification steps.

  20  

Results and Discussion

Design rationale for ProcA2.8 mutant peptides

In a previous study, Dr. Christopher Thibodeaux developed a kinetic model for the wild-

type ProcA2.8/ProcM reaction (23). In the hopes of making more precise measurements for each

individual modification step, as well as to ascertain the effects of PTMs on the rates of these

modifications, I have designed a series of ProcA2.8 variant peptides (Table 2, Figure 6), which

will be subjected to kinetic studies. In these seven ProcA2.8 mutant peptides, the Ser and/or Cys

residues involved in the modifications are mutated into Ala residues in a manner, such that only

certain modifications will occur in the presence of ProcM. The hypothesis is that simplifying the

ProcM/ProcA2.8 reaction into its constituent reactions will enable us to reduce the complexity of

the kinetic model for each variant substrate into a simpler form that will provide more accurate

rate constants upon numerical simulation and global fitting. The variants will allow us to

measure each of the two dehydration rates (via the S1 and S2 substrates) and each of the two

cyclization rates (via C1S1 and S2C2) in isolation. The more complex S1S2, C1S1S2, S1S2C2,

and wt substrates will then allow us to determine the effects of the presence of different residues

or PTMs on each dehydration and cyclization reaction.

  21  

Table 2. ProcA2.8 mutant core peptide sequences. The name of each of the mutants denotes the presence of residues (highlighted in red) that are involved in ProcM-catalyzed PTMs.

ProcA2.8 peptide species Core Peptide Sequence

ProcA2.8-wt Leader*-AACHNHAPSMPPSYWEGEC

ProcA2.8-S1 Leader-AAAHNHAPSMPPAYWEGEA

ProcA2.8-S2 Leader-AAAHNHAPAMPPSYWEGEA

ProcA2.8-S1S2 Leader- AAAHNHAPSMPPSYWEGEA

ProcA2.8-C1S1 Leader- AACHNHAPSMPPAYWEGEA

ProcA2.8-S2C2 Leader- AAAHNHAPAMPPSYWEGEC

ProcA2.8-C1S1S2 Leader- AACHNHAPSMPPSYWEGAC

ProcA2.8-S1S2C2 Leader- AAAHNHAPSMPPSYWEGEC *Leader sequence: MSEEQLKAFLTKVQADTSLQEQLKIEGADVVAIAKAAGFSITTEDLNSHRQNLSDDELEGVAGG

Figure 6. Schematic representation of ProcA2.8 mutant peptides. The name of each peptide species is indicated.  

  22  

ProcA2.8 peptide purification

Each variant peptide shown in Figure 6 was cloned, expressed, purified, and processed as

described in the Methods section. Typically, roughly 10 mg of peptide was obtained after

thrombin digestion (to remove the His6 affinity tag used for purification) and HPLC. To ensure

the purity of these peptide preparations, the HPLC purified peptide samples were further

analyzed by liquid chromatography electrospray ionization mass spectrometry (LC/ESI-MS,

Figures 7 and 8). As can be seen from the total ion chromatograms (Figure 7), each peptide

eluted from the column at around 7 min, with very little variation in the elution time between the

different mutants. No other major peaks were detected, which confirmed the purity of the HPLC

purified thrombin-digested peptides. Furthermore, the mass spectra for these samples (generated

by summing the individual spectra that compose the chromatographic peaks in Figure 7) clearly

indicated that the desired peptides are responsible for the vast majority of the total signal (Figure

8).

  23  

The ESI-mass spectra shown in Figure 8 illustrate an important point in terms of how I

quantify peptide concentration in the kinetic assays. Namely, the time-of-flight (TOF) mass

analyzer on the mass spectrometer allows people to simultaneously detect multiple charge states

for each peptide species (as shown for the wt peptide in Figure 8H). TOF detectors (like all mass

analyzers) distinguish ions based on mass-to-charge ratio (m/z). The observation of several

multiply charged ions for each peptide effectively provides replicate measurements for each

Figure 7. Total ion chromatograms for the thrombin-digested wild-type ProcA2.8 and seven mutant peptides. All of the peptides eluted as a single, broad peak under the LC conditions described in the Methods.  

  24  

peptide species in each sample that is analyzed, and is integral for the accurate quantitation of

peptide concentration as discussed in more detail below. These differentially protonated peptides

are generated during the electrospray ionization (ESI) process. ESI allows molecules to acquire

charge as they transfer to the gas phase from a liquid stream. In this case, ESI-MS is coupled

with liquid chromatography and, thus, the liquid stream is the effluent from the LC column. The

sample solution is sprayed through the tip of a capillary tube, where a high voltage is applied. A

nebulizer gas (nitrogen) flows into the capillary to help spray the sample through the tip. As the

sample is sprayed through the capillary, solvent droplets form and acquire charge at the tip of the

capillary. These charged droplets then pass down a pressure gradient where the solvent

molecules evaporate, thereby increasing the charge density on the droplet surface. As the peptide

sample transfers into the gas phase from the ESI droplet, the multiple charges on the droplet

surface are transferred to the peptide. Usually, when peptide/protein samples are analyzed by

ESI-MS, the electric field is manipulated to generate positive charges on the droplets, which are

typically transferred to basic amino acid residues (such as Lys, Arg and His) as the peptide enters

the gas phase. Because the ProcA2.8 peptides contain several basic amino acid residues, they are

typically sprayed as a mixture of several multiply charged ions (usually the 6+ through the 11+

ions are observed). Formic acid (FA) present in the LC solvent is used to ensure that the positive

charge is maximized on the gas phase peptide ions. FA also helps to protonate acidic residues

(such as Asp and Glu), which will otherwise neutralize the positive charge and decrease the

overall sensitivity of MS analysis.

  25  

ProcM Purification

ProcM was first purified from the cell lysate using Ni-NTA affinity chromatography.

Nickel has the ability to coordinate with certain chemical species. When Ni2+ is charged onto the

column, it forms four coordination interactions with the nitrilotriacetic acid (NTA) moiety that is

attached to the agarose beads. When His6-ProcM was loaded onto the column, the Ni2+ trapped

Figure 8. Electrospray ionization mass spectra for thrombin-digested wild-type ProcA2.8 and mutant peptides. Each peptide species is observed in multiple charge states. The charge states are shown in the spectra for wild-type ProcA2.8 (panel H). The observed and theoretical mass of each mutant precursor peptide is shown. Panel A: ProcA2.8-S1, panel B: ProcA2.8-S2, panel C: ProcA2.8-S1S2, panel D: ProcA2.8-C1S1, panel E: ProcA2.8-S2C2; panel F: ProcA2.8-C1S1S2, panel G: ProcA2.8-S1S2C2, panel H: wt ProcA2.8.  

  26  

the recombinant protein by coordinating to the His6-tag on the N-terminus of ProcM. After the

non-specifically bound proteins had been washed off the column with buffer containing 20 mM

imidazole, buffer containing 200 mM imidazole was used to elute the desired protein. The

elevated concentrations of imidazole displace the imidazole groups of the His6-tag from the Ni-

NTA resin.

The ProcM present in crude cell lysates tends to aggregate. Therefore, after the protein

sample had been eluted from the Ni-NTA column and concentrated to approximately 3 mL, gel

filtration was used to separate the monomer fraction from the less active oligomer and aggregate

fractions. Gel filtration separates proteins on

the basis of size. In the gel filtration column,

the stationary phase is composed of beads

containing tiny pores. Smaller molecules

tend to interact more extensively with the

pores, spend more time travelling through

the beads and thus elute more slowly. Larger

molecules interact less with the pores and

thus elute more quickly. This results in the

separation of molecules based on their sizes.

From the gel filtration chromatogram (Figure 9), it can be seen that ProcM aggregates had the

largest size and eluted first, while ProcM monomer has the smallest size and eluted last. The

activities of these various gel fractions from the gel filtration column have been previously tested

and the monomer is the most active. Therefore, only the monomer fraction was collected and

Figure 9. Gel filtration column chromatogram. The peaks corresponding to ProcM aggregate, oligomer and monomer are indicated.  

  27  

used in the subsequent kinetic studies. Usually, I obtain approximately 5 mg of purified

monomer ProcM per L culture.

Validation of kinetic assays

A brief description of the kinetic assay employed in this study and its suitability for

studying LanM/LanA reactions will be discussed below. In vitro kinetic assays were performed

as described in the Methods section for the reaction of ProcM with each of the ProcA2.8 variants

listed in Table 2. At the desired time points, aliquots from the reaction mixture were quenched

and alkylated with N-ethylmaleimide (NEM). As shown in previous control experiments, NEM

reacts rapidly and selectively with the thiol side chain of Cys residues under the conditions used

(23). This alkylation step is critical for distinguishing the isobaric cyclized and uncyclized

peptide species in the reaction mixtures by mass spectrometry. Other than the cyclization steps,

the other LanM-catalyzed modifications (dehydration and phosphorylation) can be clearly

distinguished by mass spectrometry, and tandem mass spectrometry can be used to determine the

regiochemistry of the modifications. For these reasons, ESI-MS is the ideal analytical method for

interrogating the multistep reactions catalyzed by LanM enzymes.

Each time point aliquot was analyzed by top-down LC/ESI-MS, meaning that the

peptides in the samples were not proteolyzed prior to MS analysis. My reasoning behind this

decision was two fold. First, the assay work up is simplified without peptide digestion, leaving

less chance for human error to contribute to the measurement. Second, and most importantly, Dr.

Christopher Thibodeaux reasoned that keeping the leader peptide (which is unmodified by

ProcM) attached to the core peptide might help to nullify potential differences in the ESI

  28  

ionization efficiencies of the various peptide intermediates present in the sample. Indeed, the

validation experiments performed in previous studies suggest that this is the case (23).

In this assay, the relative abundances of peptide species present in the sample are

quantified from the integrated peak areas of the ions of interest as described in the Methods and

in previous literature (27, 28). For this quantitation method to be accurate, several conditions

must be met. First, there must be a linear relationship between the LC/ESI-MS signals and the

concentration of the ProcA2.8 peptide species present in the reaction aliquots. Analysis of

serially diluted peptide stock solutions and ProcM/ProcA2.8 reaction mixtures containing

mixtures of reaction intermediates allowed us to define a linear concentration range for the assay

(from 0.2 to 10 µM) and showed that each detectable intermediate varied linearly within this

range (23). Another critical requirement for the assay to function as desired is to demonstrate that

the various post-translational modifications do not drastically alter the ionization efficiency of

the intermediates. To address this issue, Dr. Christopher Thibodeaux devised assays to

demonstrate that neither NEM alkylation nor phosphorylation were altering ProcA2.8 ionization

efficiency (23). As mentioned above, the physiochemical properties of the ProcA2.8 peptides are

likely defined mainly by the large (70 residue), unmodified leader peptide, rather than by the 20

amino acid long core peptide – enabling accurate relative quantitation by top-down MS.

Before discussing each ProcA2.8 variant reaction in detail, I will provide a brief

summary of the data fitting procedure. The rate constants for all of the kinetic data reported in

this work were determined by numerical simulation of the data with Kintek Explorer (25, 26).

The fitting process was performed by Dr. Christopher Thibodeaux. For this analysis, the starting

conditions (enzyme and substrate concentration, reaction time, and kinetic model) were specified.

All of the kinetic models included reversible enzyme-substrate binding steps for each species in

  29  

the model. The binding affinity was determined previously and was assumed to be leader peptide

dependent and constant for all observable intermediates (23). Dynamic simulation was employed

to estimate starting values for the variable parameters in the fit (i.e. the rate constants for post-

translational modification). Using the specified starting conditions, kinetic model, and parameter

estimates, the data were globally fit using a least squares fitting procedure implemented in

Kintek Explorer. The data for each substrate were initially fit independently to obtain a rough

estimate for the rate constants. After this initial fitting, the data were closely inspected in order to

identify rates for certain PTMs that tended to have similar values in the fits for different

substrates. If a given rate constant (e.g. dehydration of Ser9) was found to be similar in two

separate reactions, the two rates were set equal to one another in the global fit. In this manner,

the available parameter space was reduced, which helps to produce more realistic and

constrained fits. After the initial fits were complete, the data for all eight reactions (wt ProcA2.8

and the seven variants shown in Table 2) were globally fit simultaneously to find a single set of

best fit values for the rate constants. For the global fit, all rate constants for a given PTM were

held at fixed relative ratios using the values determined by the fits of the kinetic data for

individual substrates. Thus, the final model had five variable rate constants (dehydration of Ser9,

dehydration of Ser13, cyclization of ring A, cyclization of ring B, and phosphorylation of Ser9).

Using the FitSpace Explorer suite of Kintek Explorer, he then verified that the best fit values of

each variable parameter were well constrained (i.e. each variable rate constants could only

assume a well defined set of values while still producing a good fit). The rate constants reported

in the ensuing discussion for each individual substrate are the rates derived from the global fit.

The rate constants, error estimates, and other statistics of the global fit are provided in Table 3 at

the end of the Discussion.

  30  

Kinetic analysis of ProcA2.8 variant peptides

In order to measure the rate of dehydration at Ser9 more precisely, a kinetic assay of the

ProcA2.8-S1 substrate was performed. In the S1 mutant, other than Ser9, all of the other residues

involved in the post-translational modification (Cys3, Ser13 and Cys19) were mutated into Ala

residues such that only dehydration at Ser9 could proceed. As expected, the starting material (1)

and the final product containing one Dha residue (2) were observed in the LC/ESI-MS spectra

(Figure 10C). The simulated rate constant for Ser9 dehydration was 5.9 min-1 (for error bars

based on the fit, see Table 3). In addition, a phosphorylated intermediate (3) present at low

abundance also formed at a rate of 0.41 min-1. During simulation to the model in Figure 10A,

both rate constants were allowed to vary freely to find the best fit values.

The phosphorylated peptide is an intermediate en route to dehydration (Figure 2) and

previous studies have shown that ADP must be bound to the ProcM active site in order for

phosphate elimination to occur (23, 29). However, if the phosphopeptide dissociates from the

enzyme (prior to phosphate elimination), then the bound ADP would rapidly exchange with ATP,

which is present in large molar excess in the reaction. Thus, if the phosphopeptide is released, re-

formation of the catalytically competent ProcM:phosphopeptide:ADP complex would be

difficult under the reaction conditions. The phosphorylated intermediate would be transformed

into the product eventually (e.g. as in Figure 17E), but within the time period in the kinetic assay,

the conversion is negligible. Accordingly, phosphorylation of Ser9 (1→3) was modeled as an

irreversible side reaction in the kinetic mechanism for the ProcA2.8-S1 substrate (Figure 10A).

Similarly, all phosphorylated species observed in my studies were kinetically modeled as

irreversible, dead-end species (except for the phosphorylated species in the wt ProcA2.8, which

were observed long enough to see their consumption).

  31  

Also apparent from the reaction spectra in Figure 10C is the TCEP adduct (2*) of the

dehydrated product (2). The TCEP adducts form by nucleophilic addition of the phosphine

moiety of TCEP onto the Cβ of Dha residues. The TCEP adduct likely forms during the

derivitization of the quenched time points because high TCEP concentrations are used to ensure

efficient NEM alkylation of Cys thiols. For all TCEP adducts observed in this study, the

fractional abundance of TCEP adducts (in this case 2*) were combined with the fractional

abundance of their parent species (in this case 2) and the sum of the two species were simulated

as a single entity.

Figure 10. Kinetic analysis of the ProcM/ProcA2.8-S1 reaction. A. Kinetic model for the ProcA2.8-S1 reaction. The major species include the starting material (1), final product (2), and a phosphorylated intermediate (3). Numbers over the arrows are rate constants (min-1) B. Table of the expected mass of major peptide species and the observed masses determined from the mass

  32  

spectra. C. Mass spectral time course for the ProcM/ProcA2.8-S1 reaction, showing spectra at 2, 15, and 120 min. Only the 8+ ion family is shown, but all observed multiply charged ionic states were used for quantitation of peptide fractional abundance. D. Time courses for peptide species in the ProcM/ProcA2.8-S1 reaction. The charge-state-weighted fractional abundances of peptide species were determined as described in the methods, converted to peptide concentration, and plotted vs. reaction time. The progress curves were generated by numerical simulation of the data with the kinetic model shown in panel A. by Dr. Christopher Thibodeaux.

The kinetic analysis of ProcA2.8-S2 (Figure 11) revealed a slightly faster dehydration

rate for Ser13 (7.7 min-1) than the Ser9 dehydration rate in the ProcA2.8-S1 peptide (5.9 min-1).

This slight difference is consistent with previous studies using wt ProcA2.8 analogues deuterated

at Cα, which showed preferential dehydration of Ser13 prior to dehydration of Ser9 (24).

Notably, unlike modification of the S1 substrate, no phosphorylated intermediates were detected

in the ProcA2.8-S2 reaction, suggesting more efficient coupling of the phosphate transfer and

elimination steps at the Ser13 site. Together, these observations suggest that the core peptide in

the vicinity of Ser13 may have a slightly higher binding affinity for the dehydratase active site

than the region of the core peptide containing Ser9. In this scenario, Ser13 would bind

preferentially to the dehydratase and get modified first, and the phosphopeptide intermediate

would be less likely to dissociate prior to phosphate elimination.

  33  

Figure 11. Kinetic analysis of the ProcM/ProcA2.8-S2 reaction. A. Kinetic model for the ProcA2.8-S2 reaction, including major peptide species detected in the ESI mass spectra: starting material (4) and final product (5). B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectral time courses for the ProcM/ProcA2.8-S2 reaction recovered at 2, 8, and 240 min (the 8+ ion families is shown). D. Simulated kinetic data performed by Dr. Christopher Thibodeaux.

I next analyzed the kinetic data for the ProcA2.8-S1S2 substrate, which contains both

dehydratable residues (Ser9 and Ser13), but lacks both Cys residues and, thus, cannot be cyclized

by ProcM (Figure 12). Because the dehydration rates at Ser9 and Ser13 were not found to differ

drastically (5.9 min-1 and 7.7 min-1, respectively), either dehydration could conceivably occur

first during the maturation of ProcA2.8-S1S2. Thus, even though the data does not resolve

between the isomers having only Dha9 or Dha13 (7 and 8), I assumed that both were present in

the reaction mixture. This assumption required a kinetic model with two reciprocal pathways for

the conversion of starting material to the doubly dehydrated final product (either 6 → 7 → 9 or 6

→ 8 → 9). During simulation, the rate constants for the initial dehydration in both pathways (6

  34  

→ 7 and 6 → 8) were set equal to the corresponding dehydration rate constants determined for

the simplified S1 and S2 mutants (5.9 and 7.7 min-1, respectively). During the fit, the ratio

between these two rates was held fixed, but the absolute magnitude of the rates were allowed to

vary. The remaining parameters in the model were allowed to vary freely during the fit. This fit

resulted in nearly identical rates for the initial dehydration (as compared to the dehydration rates

measured for the S1 and S2 substrates). Interestingly, along both dehydration pathways, the

second dehydration was found to be substantially faster, consistent with the low level

accumulation of species 7 and 8 (the red curve in Figure 12D). This model suggests that the

dehydration rates of both Ser9 and Ser13 are enhanced in the presence of another Dha residue.

The most rational explanation for this effect is that the dehydratated residue may stabilize

docking interactions between the core peptide and the dehydratase. The relative ratios of the four

dehydration rate constants determined in the S1S2 fit were used in the initial fitting of all

substrates that contained both Ser residues. Thus, the models assume that either Ser can be

dehydrated first and that the dehydration rate for each site is enhanced if the other site has

already been dehydrated.

The previous experiments with the S1 and S2 analogues (as well as unpublished tandem

MS data on the wt ProcA2.8 reaction) suggested that phosphorylated ProcA2.8 intermediates

always contain the phosphate at Ser9. Therefore, although the current data does not resolve the

regiochemistry of the phosphorylations, species 10 and 11 were reasonably presumed to be the

phosphorylated versions of compounds 6 and 8, respectively. During the fit of the S1S2 data, the

rate constant for the phosphorylation of starting material (6 → 10, 0.41 min-1) was found to be

very similar to the same reaction in the S1 substrate. Thus, the two rates were held at equal

values in all subsequent fits. The rate for phosphorylation of the once-dehydrated intermediate (8

  35  

→ 11, 2.0 min-1) was faster. Thus, dehydration of Ser13 appears to enhance both

phosphorylation and net dehydration of Ser9.

Figure 12. Kinetic analysis of the ProcM/ProcA2.8-S1S2 reaction. A. Kinetic model for the ProcA2.8-S1S2 reaction. The major peptide species are the starting material (6), mono-dehydrated intermediates (7, 8), phosphorylated intermediates (10, 11) and final product (9) B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectra for the ProcM/ProcA2.8-S1S2 reaction at 2, 30, min and 240 min (the 8+ ion families is shown). D. Simulated kinetic data performed by Dr. Christopher Thibodeaux. The singly dehydrated species 7 and 8 are isobaric and were simulated as a single entity. The peak areas for the oxidized form of the starting material (6•) and the TCEP adduct (9*) were summed with 6 and 9, respectively, for fitting.

The ProcA2.8-C1S1 mutant peptide contains both Cys3 and Ser9. ProcM is expected to

facilitate ring A formation in this substrate by dehydrating Ser9, followed by establishing ring A

  36  

via Michael-type addition of Cys3 onto the Dha residue. As expected, the starting material (12),

the intermediate containing Dha9 (13) and the final product containing ring A (14) were all

observed in the mass spectra (Figure 13C). The data for the ProcA2.8-C1S1 substrate could be fit

using the same Ser9 dehydration and phosphorylation rate constants used in the S1 and S1S2 fits

(5.9 and 0.41 min-1, respectively). An additional step for the cyclization rate of ring A was

needed to complete the fit. Interestingly, the ring A cyclization rate from the fit of the ProcA2.8-

C1S1 data (0.59 min-1) is twice as fast as the rate-limiting ring A cyclization event in the wild-

type ProcA2.8 maturation model (0.30 min-1). In this model (Figure 17A), cyclization of ring A

occurs after the cyclization of ring B (23). This observation is consistent with my previous

conclusion that the presence of ring B adversely affects the rate of ring A cyclization, and that

the ProcM cyclase suffers from losses in catalytic efficiency when structure (in the form of

thioether rings) is introduced into the maturing peptide.

  37  

Figure 13. Kinetic analysis of the ProcM/ProcA2.8-C1S1 reaction. A. Kinetic model for the ProcA2.8-C1S1 reaction. The major peptide species are starting material (12), dehydrated intermediate (13), phosphorylated intermediate (15) and final product (14) B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectral time course for the ProcM/ProcA2.8-C1S1 reaction at 2, 30, and 240 min (the 8+ ion families is shown). D. Simulated kinetic data carried out by Dr. Christopher Thibodeaux.

To analyze the kinetics of ring B formation, I investigated the ProcA2.8-S2C2 substrate,

which contains only the Ser13 and Cys19 residues (Figure 14). Maturation proceeded with

precursor dehydration at Ser13 (16 → 17), followed by formation of ring B between the Dha13

and Cys19 residue (17 → 18). Once again, no phosphorylated intermediate was observed,

providing more indirect evidence that the phosphorylated species seen with the S1, C1S1, and

S1S2 substrates harbors the phosphate moiety on Ser9. Finally, as can be see in Figure 14D, the

consumption of the starting material (16) was biphasic, suggesting a two-step process. This

  38  

behavior was seen previously with the wt ProcA2.8 and was attributed to the presence of an

inactive form of substrate that is slowly converted to the active form (23). Thus, the kinetic

model shown in Figure 14A included an addition step (16* → 16, Table 3) where the inactive

starting material is converted to 16 at a rate of 0.005 min-1. The inactive substrate likely involves

formation of peptide aggregates via intermolecular disulfide bonds (23).

During the simulation process, the initial value for the Ser13 dehydration rate constant

(16 → 17) was set to be the same as the Ser13 dehydration rate determined for the S2 and S1S2

substrates (7.7 min-1) and the cyclization rate (17 → 18) was adjusted to give simulated curves

that best approximated the experimental data. After fitting, the Ser13 dehydration rate was

indeed very similar to the rates determined from the S2 and S1S2 substrates. The ring B

cyclization rate determined from this fit (1.2 min-1) is somewhat faster than the ring A

cyclization rate (0.59 min-1) determined for the C1S1 peptide, perhaps providing a kinetic basis

for the previously observed preferred order of cyclization (namely, ring B forms first in the wt

substrate, followed by ring A) (24).

  39  

Figure 14. Kinetic analysis of the ProcM/ProcA2.8-S2C2 reaction. A. Kinetic model for the ProcA2.8-S2C2 reaction. Major species include starting material (16), dehydrated intermediate (17), and final product (18). B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectral time course for the ProcM/ProcA2.8-S2C2 reaction at 2, 20, and 240 min (the 8+ ion family is shown). D. Simulated kinetic data carried out by Dr. Christopher Thibodeaux. The oxidized species (16• and 18•) were combined with their parent ions (16 and 18, respectively) for fitting. In addition, the unalkylated starting material (16°) seen in some of the samples was combined with the signals for 16 and 16•.

I next turned my attention to the more complicated ProcA2.8 substrates, C1S1S2 and

S1S2C2 (Figures 15 and 16, respectively). Like the S1S2 substrate, the kinetic models for both

the C1S1S2 and S1S2C2 substrates included a branched dehydration pathway, where the first

dehydration could occur either at Ser13 or at Ser9. During simulation of the individual data sets,

the relative rates of the four dehydration steps in the ProcA2.8-C1S1S2 and ProcA2.8-S1S2C2

models were held identical to the relative rates of the dehydration steps in the S1S2 substrate.

Interestingly, the dehydration rates for both the C1S1S2 and S1S2C2 substrates had to be

substantially slowed (roughly 5 fold) in order to achieve good fits. Thus, increased complexity of

  40  

the ProcA2.8 core peptide seems to correlate with slower dehydration rates. Perhaps the Cys and

Ser residues help the core peptide to engage in interactions with ProcM that inhibit access of the

ProcM dehydratase active site to the modification sites of ProcA. As more of these sites are

added, the inhibitory interactions between the core peptide and ProcM get stronger while

dehydration gets slower. This may be a general property of ProcM with each of its 30 natural

substrates.

Interesting properties also emerge when looking at the cyclization rates in the C1S1S2

and S1S2C2 substrates. The cyclization rate of ring B in the S1S2C2 substrate (Figure 16A, 28

→ 29, 1.1 min-1) was nearly identical to the cyclization rate of ring B in the simplified S2C2

substrate (Figure 14A, 17 → 18, 1.2 min-1), suggesting no effect of the presence of Dha9 on the

ProcM-mediated cyclization of ring B. Thus, docking of Dha13 and Cys19 into the active site of

the ProcM cyclase domain, seems to be independent of other structural features of the core

peptide. In contrast, there was a significant increase in the rate of ring A formation in the

C1S1S2 substrate (1.5 min-1) relative to the C1S1 substrate (0.59 min-1). This suggests that,

similar to Ser9 phosphorylation and dehydration, ring A cyclization is somehow enhanced by the

presence of Dha13, most likely through stabilizing binding interactions of the two-fold

dehydrated C1S1S2 species (22, Figure 15A) to the cyclase active site of ProcM. These

differential effects of peptide structure on the cyclization rates of rings A and B suggest that

differing molecular interaction between the core peptide and the ProcM cyclase could be what

determines the rates of thioether ring formation mediated by ProcM.

  41  

Figure 15. Kinetic analysis of the ProcM/ProcA2.8-C1S1S2 reaction. A. Kinetic model for the ProcA2.8-C1S1S2 reaction. The major species include the starting material (19), mono-dehydrated intermediates (20, 21), double-dehydrated intermediate (22), phosphorylated intermediate (24) and the final product (23). B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectral time course for the ProcM/ProcA2.8-C1S1S2 showing the 2, 30, and 240 min time points (the 8+ ion families is shown). D. Simulated kinetic data carried out by Dr. Christopher Thibodeaux.

  42  

Figure 16. Kinetic analysis of the ProcM/ProcA2.8-S1S2C2 reaction. A. Kinetic model for the ProcA2.8- S1S2C2 reaction. The major peptide species include starting material (25), mono-dehydrated intermediates (26, 27), double-dehydrated intermediate (28), phosphorylated intermediate (30) and the final product (29). B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectral time course for the ProcM/ProcA2.8-S1S2C2 reaction, showing the 2, 30, and 240 min time points (the 8+ ion families is shown). D. Simulated kinetic data carried out by Dr. Christopher Thibodeaux.

Kinetic analysis of wt ProcA2.8 and summary of kinetic trends

In light of the kinetic analysis of the ProcA2.8 variants presented above, the published

kinetic data for the wild-type ProcA2.8 peptide were refit with the slightly modified kinetic

model shown in Figure 17A. In the previous study, our laboratory had not yet made independent

measurements of the dehydration rates of Ser9 and Ser13. Thus, the data was conservatively fit

  43  

with a model where Ser13 dehydration preceded Ser9 dehydration in a linear reaction sequence

(23). Here, we simply show that the wt data can be satisfactorily refit with the branched

dehydration mechanism and relative rate ratios employed in the S1S2, C1S1S2 and S1S2C2

models discussed above (Figure 10-16). This re-fit led to only minor changes in the fitted values

for the other rate constants in the model, with the most drastic effect being the reduction in the

value of the second dehydration step from 13 min-1 in the original model to either 2.7 or 4.5 min-

1 depending on the order of dehydration.

The trends in the cyclization rates suggest a relationship between peptide structure and

cyclization rate. The cyclization rate of ring A seems to be modulated by several factors. First,

the presence of ring B seems to drastically slow ring A formation. Previous studies using tandem

mass spectrometry and chemically synthesized ProcA2.8 analogues have shown that ring B

formation precedes ring A formation in the wt ProcA2.8 peptide (24). The kinetic studies

reported here of the wt peptide show that the first ring (ring B) forms at a rate of 2 min-1, while

ring A forms an order of magnitude more slowly (0.3 min-1). In the C1S1 and C1S1S2 peptides,

the ring A cyclization rate increases to 0.59 min-1 and 1.5 min-1, respectively, clearly revealing

the inhibitory effect of ring B on ring A formation kinetics. On the basis of the kinetic data for

the wt substrate, our group previously proposed that the ProcM cyclase domain does not tolerate

the more defined structure of partially cyclized the core peptides (23). The current data (in

addition to unpublished kinetic data on other ProcM substrates) corroborates this hypothesis.

Also, the data reported here show that ring A cyclization is enhanced (form 0.59 min-1 to 1.5

min-1) when both Ser9 and Ser13 are dehydrated. Given the similarity in the cyclization rate of

ring A in the C1S1S2 substrate (1.5 min-1) to the cyclization rate of ring B in the wt substrate

(2.0 min-1), it is unclear why there is such a preference for cyclizing ring B first in the wt

  44  

substrate. Perhaps there is preferential coordination of Cys19 to the Zn ion the cyclase active site,

which poises the enzyme for ring B cyclization in the context of the wt substrate. Compared

with ring A, the cyclization rate of ring B seems to be relatively insensitive to the core peptide

structure. Namely, the ring B cyclization rates followed the order wt (2.0 min-1) > S1S2C2 (1.1

min-1) ~ S2C2 (1.2 min-1). Thus, the rate of ring B cyclization may be more strongly governed by

its C-terminal positioning on the core peptide, rather than by the local amino acid sequence

context.

  45  

  46  

Figure 17. Kinetic analysis of the ProcM/wt ProcA2.8 reaction. A. Kinetic model for the wt ProcA2.8 reaction. The major peptide species include starting material (31), mono-dehydrated intermediates (32, 33), double-dehydrated intermediate (34), cyclized intermediate with one ring (35), phosphorylated intermediates (37-39) and the final product (36). B. Table of the expected mass of major peptide species and the observed masses determined from the mass spectra. C. Mass spectral time course for the ProcM/ProcA2.8 reaction showing the 3, 90, and 1350 min time points (the 8+ ion families is shown). D and E. Simulated kinetic data. Table 3. Summary of simulated parameters for the global fit of the kinetic dataa

Binding Rateb reaction ratec boundaryd 𝑘!" 4.2 - 𝑘!"" 20 -

ProcA2.8-S1 reaction rate boundary 1 → 2e 5.9 4.7 – 7.4 1 → 3 0.41 0.26 – 0.59

ProcA2.8-S2 reaction rate boundary

4 → 5 7.7 6.2 - 10 ProcA2.8-S1S2

reaction rate boundary 6 → 7 5.9 4.7 – 7.4 6 → 8 7.7 6.2 - 10 7 → 9 25 20 - 33 8 → 9 19 15 - 24

6 → 10 0.41 0.26 – 0.59 8 → 11 2.1 1.3 – 3.0

ProcA2.8-C1S1 reaction rate boundary 12 → 13 5.9 4.7 – 7.4 13 → 14 0.55 0.41 – 0.60 12 → 15 0.41 0.26 – 0.59

ProcA2.8-S2C2 reaction rate boundary 16 → 17 7.7 6.2 - 10 17 → 18 1.1 0.74 – 1.8

16* → 16 0.005 -

ProcA2.8-C1S1S2 reaction rate boundary 19 → 20 1.2 0.98 – 1.5 19 → 21 2.0 1.6 – 2.6 20 → 22 5.1 4.1 – 6.6 21 → 22 4.9 3.9 – 6.1 22 → 23 1.5 1.1 – 1.9 19 → 24 0.15 0.1 – 0.22

ProcA2.8-S1S2C2 reaction rate boundary 25 → 26 1.2 0.98 – 1.5 25 → 27 2.0 4.1 – 6.6 26 → 28 5.1 1.6 – 2.6 27 → 28 4.9 3.9 – 6.1 28 → 29 1.1 0.74 – 1.8 27 → 30 0.55 0.35 – 0.80

wt ProcA2.8 reaction rate boundary 31 → 32 0.85 0.68 – 1.06 31 → 33 1.4 1.1 – 1.8 32 → 34 4.5 3.6 – 5.9 33 → 34 2.7 2.1 – 3.3 34 → 35 1.96 1.3 – 3.2 35 → 36 0.30 0.23 – 0.38 31 → 37 0.14 0.09 – 0.20 37 → 38 0.15 0.12 – 0.19 38 → 39 >> 0.15 - 39 → 35 0.08 0.05 – 0.12

31* → 31 0.005 - a Statistics of global fit: χ2 = 2429, N = 914, DOF = 909, χ2/DOF = 2.7 b Binding rate constants were assumed to be the same for each intermediate (1 – 39). See ref 23. c 𝑘!" is the bimolecular rate constant for formation of the enzyme:substrate Michaelis complex and is in units of µM-1min-1. All other rate constants are in units of min-1.    𝑘!"" is the dissociation rate of the Michaelis complex. d Boundaries were calculated by FitSpace at 1.1 times the χ2 minimum. See refs 23, and 25-26. e Rate constants are color-coded by the reaction type and by how they were linked during the global fit: black – held constant; red: Ser9 dehydration; green – Ser13 dehydration; gold – ring A formation; purple – ring B formation; blue: Ser9 phosphorylation.

  47  

Conclusion:

In summary, in the hopes of understanding how ProcM maintains its relaxed substrate

specificity, the post-translational modification of ProcA2.8 analogs catalyzed by ProcM has been

characterized in this work. Previous kinetic studies on the ProcM/ProcA2.8 reaction suggested

that conformational changes and protein-protein interactions between ProcM and ProcA2.8

substrates may affect the relative rates and, hence, the order of PTMs. In other words, during

maturation, the conformation of the ProcA precursor peptides gradually change, thereby altering

their interactions with ProcM, which subsequently affects the kinetic parameters of the ProcM-

installed PTMs. In an attempt to provide direct experimental evidence for this hypothesis, a

series of ProcA2.8 variants were constructed and the kinetics of their modification was

investigated.

As described above, the rate constants for certain PTMs do indeed seem to be altered by

the structure and PTM state of the precursor peptide. Most notably, the rate of ring A cyclization

(which is the slowest and final reaction in the maturation of the wt peptide) is substantially faster

in the absence of ring B – reaching a rate in the C1S1S2 variant (1.5 min-1) that rivals the rate of

ring B formation in the wt peptide (2.0 min-1). This data suggests that there is nothing

intrinsically difficult in the molecular recognition of the residues that form ring A by ProcM.

Rather, it seems to be the presence of the ring B thioether structure in the wt peptide that

adversely affects the ring A cyclization rate. Perhaps the steric constraints induced by ring B

formation limit the accessibility of the residues that form ring A by the ProcM cyclase. Indeed,

Ser9 and Ser13 are separated by the sequence MetProPro. In conjunction with ring B, the double

Pro motif may place severe restrictions on the conformation of the core peptide sequence and

limit the conformational space available for sampling by the enzyme:substrate Michaelis

  48  

complex. The rules that govern ring topology in the prochlorosins are still largely unclear, but

the present work hints at an integral role for the primary amino acid sequence and the

accessibility of peptide binding by the cyclase as key factors in cyclization kinetics. Additional

studies on other prochlorosins should help to better illuminate these rules.

Another interesting observation is the change in the dehydration rates as a function of

core peptide structure. Two major trends emerge. First, dehydrations get faster if the other

residue is already dehydrated. Second, as the peptide structure gets more complex, the

dehydration rates decrease significantly. Both trends seem to be consistent with kinetics

governed by binding interactions between the peptide and the synthetase. Namely, Dha residues

may stabilize binding of the core to the dehydratase, enabling more facile dehydration of other

sites. Similarly, the presence of Ser and Cys residues on the core peptide could allow the peptide

to engage in binding interactions with the synthetase in a manner that prevents the dehydratase

domain from productive binding to its modification sites. These findings suggest that a complex

network of protein-protein interaction may be instrumental in guiding the PTM path for a given

ProcA precursor.

In summary, by revealing effects of core peptide structure on maturation kinetics, the

kinetic study accomplished in this thesis provides additional evidence in support of a previously

proposed model for ProcM function (23). This model invokes peptide-protein binding

interactions as the key modulators of the kinetic properties of the system. The generality of this

mechanism must await characterization of other prochlorosins and other lanthipeptide

biosynthetic systems. Understanding the detailed functional mechanisms of lanthipeptide

synthetases may enable the manipulation of these enzyme systems for the engineering of new

antibiotic therapeutics.

  49  

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