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Theses and Dissertations in Biochemistry Biochemistry, Department of
Spring 4-15-2020
THE ROLE OF CONFORMATIONAL DYNAMICS IN ISOCYANIDE HYDRATASE CATALYSIS
Medhanjali Dasgupta University of Nebraska - Lincoln, [email protected]
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Dasgupta, Medhanjali, "THE ROLE OF CONFORMATIONAL DYNAMICS IN ISOCYANIDE HYDRATASE CATALYSIS" (2020). Theses and Dissertations in Biochemistry. 29. https://digitalcommons.unl.edu/biochemdiss/29
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THE ROLE OF CONFORMATIONAL DYNAMICS IN ISOCYANIDE HYDRATASE CATALYSIS
by
Medhanjali Dasgupta
A DISSERTATION
Presented to the Faculty of
The Graduate College at the University of Nebraska
In partial Fulfillment of Requirements
For the Degree of Doctor of Philosophy
Major: Biochemistry
Under the Supervision of Professor Mark A. Wilson
Lincoln, Nebraska
May 2020
THE ROLE OF CONFORMATIONAL DYNAMICS IN ISOCYANIDE HYDRATASE CATALYSIS
Medhanjali Dasgupta, Ph.D.
University of Nebraska, 2020
Advisor: Dr. Mark. A. Wilson
Post-translational modification of cysteine residues can regulate protein
function and is essential for catalysis by cysteine-dependent enzymes. Covalent
modifications neutralize charge on the reactive cysteine thiolate anion and thus
alter the active site electrostatic environment. Although a vast number of enzymes
rely on cysteine modification for function, precisely how altered structural and
electrostatic states of cysteine affect protein dynamics, which in turn, affects
catalysis, remains poorly understood.
Here we use X-ray crystallography, computer simulations, site directed
mutagenesis and enzyme kinetics to characterize how covalent modification of the
active site cysteine residue in the enzyme, isocyanide hydratase (ICH), affects the
protein conformational ensemble during catalysis. Our results suggest that
cysteine modification may be a common and likely underreported means for
regulating protein conformational dynamics. This thesis will also include ongoing
work with ICH homologs, showing that Cys covalent modification-gated helical
dynamics in Cys dependent enzymes is common to enzymes of this family.
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Copyright: Medhanjali Dasgupta iii
Dedication
To my wonderful husband, Karl Francis Joseph,
&
To my emotional support cat, Scoopy Noopers.
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Acknowledgements
I would like to extend my gratitude towards my advisor, Dr. Mark A. Wilson for his support and guidance throughout my entire doctoral studies. I would also like to take this opportunity to thank my wonderful committee members, Drs. Melanie
Simpson, Joe Barycki, Donald Becker, David Berkowitz for their valuable mentoring, guidance, support and help. Of course, none of my learning would be complete without my helpful lab mates, Madison, Lin, Nate, Peter and Ekta. I also thank the collaborators who made all this work possible, Henry van den Bedem
(SSRL), Mike Wall (LANL), Aina Cohen (SSRL), Ray Sierra (LCLS), Darya
Marchany Rivera, Ken Nickerson (UNL), Javier Seravalli (UNL), Virendra
Tiwari (UNL), Dominik Budday, Saulo de Oliveira, Brandon Hayes, Sebastian
Boutet, Mark Hunter, Ruberto Alonso Mori, Alexander Batyuk, Jennifer
Wierman, Artem Lyubimov, Aaron Brewster, Nicholas Sauter, Gregory
Applegate, Michael Thompson, and James Fraser.
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Funding
This work was supported by Nebraska Tobacco Settlement Biomedical Research
Development Fund. Use of the Stanford Synchrotron Radiation Lightsource, SLAC
National Accelerator Laboratory, was supported by the U.S. Department of Energy,
Office of Science, Office of Basic Energy Sciences under Contract No. DE-AC02-
76SF00515. The SSRL Structural Molecular Biology Program is supported by the DOE
Office of Biological and Environmental Research, and by the National Institutes of
Health, National Institute of General Medical Sciences (including P41GM103393). Use of the Linac Coherent Light Source (LCLS), SLAC National Accelerator Laboratory, is supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy
Sciences under Contract No. DE-AC02-76SF00515. This research used resources of the
Advanced Photon Source, a U.S. Department of Energy (DOE) Office of Science User
Facility operated for the DOE Office of Science by Argonne National Laboratory under
Contract No. DE-AC02-06CH11357. Use of BioCARS was also supported by the
National Institute of General Medical Sciences of the National Institutes of Health under grant number R24GM111072.
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Table of Contents
CHAPTER 1: Introduction: Isocyanides as natural products.
1.1.1 Terrestrial isocyanides …………………… 3 1.1.1.a Biosynthesis of terrestrial isocyanides …………………… 3 1. 1.1.b Identification of IsnA/B homolog gene clusters …………………… 5 1.1.1.c Biological targets of terrestrial isocyanides …………………… 13 1.1.1.d Types of terrestrial isocyanides …………………… 19 Xanthocillin type …………………… 19
Cyclopentyl type …………………… 25 Assorted others …………………… 30 1.1.2 Non terrestrial isocyanides …………………… 32 1.1.2 A Types of non terrestrial isocyanides …………………… 32
Monoterpenoid type …………………… 33 Diterpenoid type …………………… 37 Sesquiterpenoid type …………………… 38 1.1.3 Degradation of isocyanides via Isocyanide hydratase …………………… 42 (ICH)
CHAPTER 2: Introduction: The relationship between internal enzyme conformational dynamics and catalysis
2.1 How internal protein motions effect the catalytic mechanism …………………… 45 of enzyme action
CHAPTER 3: Introduction: Biochemistry of reactive cysteine residues in enzymes.
3.1 The relationship between cysteine covalent modification ………………… 52 gated enzyme dynamics and catalysis.
3.2 Cysteine covalent modification and internal enzyme ………………… 53 dynamics
3.3. Cysteine covalent modification and internal enzyme ………………… 64 dynamics
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CHAPTER 4: Mix and Inject Serial Femtosecond Crystallography at XFEL reveals gated conformational dynamics during ICH catalysis.
4.1 Abstract ………………… 68 4.2 Introduction ………………… 69 4.3 Materials and Methods ………………… 71 4.4 Results ………………… 81 4.5 Discussion ………………… 96
CHAPTER 5: Purification, Kinetics and Structural Characterization of novel ICH homolog, RsICH from Ralstonia solanacearum GMI1000.
5.1 Abstract ………………… 102 5.2 Introduction ………………… 103 5.3 Materials and Methods ………………… 104 5.4 Results ………………… 110 5.5 Discussion ………………… 119
Works cited. ………………… 120
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List of Tables
Table 1: Pseudomonas ICH Enzyme kinetic parameters ……………… ix
Table 2: Pseudomonas ICH Crystallographic Data Statistics ……………… x
Table 3: Pseudomonas ICH crystallographic Refinement Statistics ……………… xi
Table 4: Ralstonia ICH Enzyme kinetic parameters ……………… xii Table 5: Ralstonia ICH Crystallographic Refinement Statistics ……………… xiii
Table 6: Ralstonia ICH Crystallographic Refinement Statistics ……………… xiv
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List of Multimedia Objects
Table 1: Pseudomonas ICH Enzyme kinetic parameters
*measured at 160 μM p-NPIC and [enzyme]=50 nM.
Standard deviations are derived from the fit; data measured n≥3
PfICH Enzyme WT G150A G150T Steady State Kinetics -1 kcat (sec ) 0.248±0.031 0.025±0.002 0.046±0.003 KM (μM) 9.262±3.250 1.208 ±0.6134 8.892±1.529 -1 -1 4 4 3 kcat/KM (M sec ) 2.68x10 2.07x10 5.21x10 Stopped Flow mixing Burst rate constant (sec-1)* 11.395±0.120 11.882±0.071 4.255±0.187 Second order rate constant for 4.85x104±943 5.51x104±557 1.11x104±999 burst (M-1 sec-1) -1 * Steady state rate kobs (sec ) 0.268±0.001 0.046±0.002 0.095±0.003
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Table 2: Pseudomonas ICH Crystallographic Data Statistics
Values for the highest resolution bins are shown in parenthesis
WT ICH WT ICH G150A G150T WT ICH WT ICH WT ICH WT ICH RT1 RT1 (more ICH ICH XFEL: XFEL: 15 XFEL: 5 cryo Sample (less oxidized) Apo1 sec min1 oxidized) Thioimida te1 Diffraction SSRL SSRL SSRL SSRL LCLS LCLS LCLS APS source 12-2 7-1 12-2 7-1 MFX MFX MFX 14BM-C Wavelength 0.827 0.975 0.827 0.975 1.305 1.305 1.305 0.900 (Å) Temperature 274 277 274 277 298 298 298 100 (K) Rayonix Rayonix Rayonix ADSC ADSC ADSC Detector Pilatus 6M Pilatus 6M MX170- MX170- MX170- Q315 Q315 Q315 HS HS HS
Space group P21 P21 P21 C2 P21 P21 P21 P21 57.24 57.15 57.04 72.11 56.88 56.77 56.81 56.58 a, b, c (Å) 58.03 57.97 57.90 59.74 57.68 57.42 57.64 56.47 69.09 69.09 69.11 56.13 68.78 68.79 68.76 68.23 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 α, β, γ (°) 112.83 112.77 112.55 115.86 112.74 112.74 112.74 112.49 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 Mosaicity (°) 0.08 0.08 0.08 0.07 N/A5 N/A5 N/A5 0.27 Resolution 39.04-1.20 42.87-1.15 38.96-1.30 35.45-1.10 20.13-1.55 20.09-1.55 20.1-1.55 37-1.05 range (Å) (1.22-1.20) (1.17-1.15) (1.32-1.30) (1.12-1.10) (1.57-1.55) (1.57-1.55) (1.57-1.55) (1.09-1.05)
Total no. of 363112 517060 327591 224032 3032726 3185312 2648001 1077211 observations (16358) (15585) (13542) (2017) (18818) (16342) (38049) (79769)
No. of unique 126373 145885 100249 84423 60075 60053 60082 179423 observations (5922) (6657) (4568) (1216) (2983) (2970) (2988) (17341) Completeness 97.4 (92.7) 98.9 (91.2) 98.1 (90.1) 97.2 (95.9) 99.9 (99.8) 99.9 (99.4) 99.9 (100) 97.1 (94.2) (%) Multiplicity 2.9 (2.8) 3.5 (2.3) 3.3 (3.0) 2.7 (2.0) 50.5 (6.3) 53.0 (5.5) 44.1 (12.7) 6.0 (4.6) 〈I/σ(I)〉 10.4(1.8) 12.5 (0.8) 9.1 (1.2) 9.7 (1.4) 60.5 (2.0) 58.4 (1.8) 73.1 (3.4) 22.8 (2.1)
2 0.991 0.999 0.996 0.999 0.962 0.963 0.937 4 CC1/2 N/A (0.225) (0.488) (0.318) (0.690) (0.16) (0.14) (0.36)
Rmeas (Rsplit for 0.072 0.059 0.066 0.045 0.196 0.197 0.232 0.07 (0.70) XFEL data)3 (1.689) (0.959) (1.513) (0.545) (0.862) (0.905) (0.658)
1 RT, Room temperature synchrotron radiation collection; XFEL, X-ray free electron laser 2 CC1/2 (31) was used to determine the high resolution cutoff. 3 Rsplit is provided only for XFEL serial crystallographic data 4 Dataset from PDB 3NON, processed before the introduction of CC1/2 5 N/A=not applicable for serial crystallographic data
xi
Table 3: Pseudomonas ICH Crystallographic Refinement Statistics
WT ICH WT ICH G150A G150T WT WT ICH WT ICH WT ICH WT RT1 (less RT1 ICH ICH ICH XFEL: XFEL: 5 RT1 (less ICH oxidized) (more XFEL: 15 sec min1 oxidized) cryo Model oxidized) Apo1 Thioimid qFit with ate1 model helix disorde r PDB code 6NI6 6NI7 6NI5 6NI4 6NPQ 6UND 6UNF 6NI9 6NJA Temperature (K) 274 277 274 277 298 298 298 274 100 Refinement PHENIX PHENIX PHENI PHENI PHENI PHENIX PHENIX PHENIX PHENI program 1.9 1.9 X 1.9 X 1.9 X 1.9 1.9 1.9 1.9 X 1.9 Resolution range 31.84- 38.53- 33.56- 35.14- 20.13- 20.09- 17.46- 31.84- 37.82- (Å) 1.20 1.15 1.30 1.10 1.55 1.55 1.55 1.20 1.05 Completeness (%) 95.17 98.87 97.83 97.16 99.93 99.86 99.96 95.00 97.11 No. of reflections 123572 145829 99966 84275 59628 59214 59498 123572 179399 No. of reflections, 6203 7300 4970 4186 2002 1986 1995 6203 8979 test set 0.1140 0.1128 0.1192 0.1098 0.1560 0.1636 0.1727 0.1078 0.1170 Rwork (0.2281) (0.2633) (0.2745) (0.2294) (0.3075) (0.3244) (0.2777) (0.2207) (0.2105) 0.1398 0.1353 0.1519 0.1283 0.1864 0.1906 0.1918 0.1402 0.1342 Rfree (0.2611) (0.2607) (0.3159) (0.2332) (0.3244) (0.3177) (0.2836) (0.2628) (0.2487) No. of non-H atoms Protein 4486 4446 4892 1868 3475 3449 3500 6192 4551 Water 342 323 323 149 298 271 272 382 484 Total 4828 4769 5215 2017 3773 3720 3772 6574 5035 Average R.M.S. deviations Bonds (Å) 0.007 0.008 0.014 0.011 0.007 0.004 0.005 0.011 0.009 Angles (°) 0.909 1.020 1.290 1.376 0.850 0.729 0.801 1.399 1.111 2 Average B factors (
Values for the highest resoltions bin shown in parenthesis 1 RT, Room temperature synchrotron radiation collection; XFEL, X-ray free electron laser 2 Anisotropy is defined as the ratio of the smallest to largest eigenvalue of the ADP tensor
xii
Table 4: Ralstonia ICH Enzyme kinetic parameters
Standard deviations are derived from the fit; data measured n≥3 RsICH Enzyme WTRsICH E122D E122Q E122T Steady State Kinetics -1 kcat (sec ) 75.65 0.15 0.37 0.34 KM (μM) 116.9 1.28 6.79 27.59 -1 -1 kcat/KM (μM sec ) 0.65 0.12 0.05 0.01 Stopped Flow mixing* Pre steady state rate 0.11274 11.882±0.071 4.255±0.187 0.16851 constant (sec 1)
*measured at 80 μM p-NPIC
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Table 5: Ralstonia ICH Crystallographic Refinement Statistics Data collected at University of Nebraska Macromolecular Structural Core Facility Model WT RsICH WT WT RsICH E122T E122D (plate) RsICH (tetragonal) (plate) (plate) (rod) PDB code NA NA NA NA NA Temperature (K) 110 110 110 110 110 Detector Raxis IV++ Raxis Raxis IV++ Raxis Raxis IV++ IV++ IV++ Refinement Refmac5 Refmac5 Refmac5 Refmac5 Refmac5 program (CCP4) (CCP4) (CCP4) (CCP4) (CCP4) Resolution range 40.21-1.50 38.52-1.46 44.9-1.63 38.42- 33.46-1.46 (Å) 1.46 Completeness 100.0 94.7 100.0 96.0 99.5 (%)(overall) Space Group P 21 21 21 P 21 21 21 P 41 21 2 P 21 21 P 21 21 21 21 a, b, c (Å) 47.84 48.13 63.50 47.90 48.20 89.17 89.04 63.50 89.19 89.20 93.04 92.77 121.46 92.88 92.96 α, β, γ (°) 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 11.0 (1.1) 13.6 (3.5) 21.0 (4.3) 26.8 22.5 (5.5) 〈I/σ(I)〉 (4.7) 0.997 0.995 0.999 0.998 0..999 CC1/2 (overall) (0.581) (0.891) (0.97) (.949) (0.952) No. of unique 65123 65535 31929 66459 69583 observations No. of 362961 341308 394901 888305 817104 observations (overall) Rfree 0.1975 0.1812 0.2002 0.1931 0.19308
Rmerge 0.73 0.66 0.62 .057 .066 (0.34) (0.926) (0.302) (0.308) (0.528)
MolProbity 2.45 2.25 2.82 2.84 4.75 clashscore Ramachandran 97.95 97.95 96.8 97.72 97.49 Favored (%) Ramachandran 0.46 0.91 1.37 0.68 1.14 Outliers (%)
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Table 6: Ralstonia ICH Crystallographic Refinement Statistics Data collected at SSRL beamline 12-2 from single crystal with 0.2°/ image and 0.15 sec/image Model WT RsICH E122D E122Q E122T
PDB code NA NA NA NA Temperature 100K 100K 100K 100K (K) Detector Pilatus 6M Pixel Pilatus 6M Pixel Pilatus 6M Pixel Pilatus 6M Pixel Array Detector Array Detector Array Detector Array Detector (PAD) (PAD) (PAD) (PAD) Wavelength 0.729 0.729 0.729 0.729 (Å) Attenuation 96% 96% 96% 96% (%) Refinement XDS, Aimless, XDS, Aimless, XDS, Aimless, XDS, Aimless, program CCP4 (Refmac 5) CCP4 (Refmac 5) CCP4 (Refmac 5) CCP4 (Refmac 5) Resolution 38.58 (0.74) 38.6 (0.82) * 38.51 (0.95) range (Å) Completenes 98.0 (89.5) 99.8 (97.4) * 99.8 (98.7) s (%)(overall) Space Group P 21 21 21 P 21 21 21 P 21 21 21 P 21 21 21 a, b, c (Å) 48.23, 89.15, 92.81 48.22, 89.2, 93.01 * 48.03, 89.39, 92.93
α, β, γ (°) 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00 90.00
19.7 (0.9) 20.1 (1) * 14.6 (1.2) 〈I/σ(I)〉 1.0 (0.322) 1.0 (0.346) * 0.999 (0.492) CC1/2 (overall) Mosaicity 0.06 0.09 * 0.05
No. of 517136 389097 * 250613 unique observations No. of 3554784 3315656 * 2754808 observations (overall) Rfree 0.1259 0.1155 * 0.13364
Values for the highest resolution bin shown in parenthesis *pending refinement
1
Chapter 1: Introduction Isocyanides as natural products
Isocyanides, also referred to as isonitriles or carbylamines, are isomers of cyanides (-C≡N). Isocyanide is the IUPAC approved nomenclature for these compounds, although the word “isonitriles” is still widely used, but not preferred. Isocyanides possess the functional group N≡C, with the organic moiety connected to the isocyano group (N≡C) via the N atom. The discovery of this unusual class of compounds was an accident; in 1859 Lieke 1 reacted allyl iodide with silver cyanide, intending to produce cyanide, but instead, ended up with a foul-smelling compound that converted to formic acid upon acid hydrolysis2. This particularly vile smelling product was recognized as a member of a new class of compounds, termed “Isomeric to Nitrile”, or “Isonitrile”3. The newly discovered isocyanides were unique in that they were the only stable organic entities whose carbon was bonded to a single atom.
Fig. 1: Zwitterionic and carbene representations of the isocyano group.
Isocyanides are highly reactive; the carbon atom of the isocyano group possesses both electrophilic as well as nucleophilic character. This makes isocyanides ideal for synthetic applications4, such as multicomponent condensation reactions5,6,7, carbohydrate and co- ordination chemistry8,9,10, addition reactions11,12 etc. Several studies have been performed to understand their electronic and geometric structures in order to explain their dual electrophilic and nucleophilic characters. For a while, the two most widely accepted forms of isocyanides were the carbenic form 13 and the zwitterionic form14 (Fig. 1). 2
These, however, are not without their individual caveats; the carbenic form was not consistent with the dipole moment and the IR- C-N stretching band at 2130 cm-115, whereas the zwitterionic form could not explain the electrophilic character of isocyanides. This structural disparity was finally addressed by performing Valence bond calculations, which supported a predominantly carbenic structure for isocyanides, with the zwitterionic resonance form being a secondary contributor. The unusual isocyano group allows isocyanides to function as biologically active naturally derived compounds such as chelators of transition metals, antibiotics, antiviral agents, marine sponge derived metabolites etc16.
In 1948, Rothe 17 isolated the first naturally occurring isocyanide, Xanthocillin, from the fungus Penicillium notatum. Xanthocillin was structurally characterized by
Hagedorn and Tonjes 18,19 in 1957, almost 100 years after Lieke’s “accidental” chemical synthesis of the first allyl isocyanides. The first marine-type isocyanide to be isolated was axisonitrile-1, from the marine sponge Axinella cannabina 20. Since then, a multitude of isocyanides and related compounds have been isolated from both marine and non-marine sources. The division of isocyanide natural products into terrestrial and marine clades has its roots in the history of their discovery as well as differences in their biosynthetic precursors, where known. This thesis will adhere to this overarching classification scheme for isocyanide natural products, although we note that other chemical taxonomies for these compounds are possible and perhaps desirable. Here, we will cover the biochemistry of isocyanide natural products, including their biological targets, biosynthetic and degradative pathways in marine vs non-marine sources as well as identifying areas where open questions invite future inquiry. 3
1.1.1 Terrestrial isocyanides: Many isocyanides have been isolated from non-marine microbial sources. Terrestrial isocyanides can be classified into three primary classes based on their structures: xanthocillins, cyclopentyls and assorted others 2,19.
1.1.1.a Biosynthesis of terrestrial isocyanides: Isocyanides are naturally produced in certain microbes as secondary metabolites. Remarkably, fundamental questions remain about the installation of the isocyanide moiety, including the still-unknown mechanisms of most validated isocyanide biosynthetic enzymes.
Isocyanides, as mentioned previously, possess a unique functional group and are naturally produced in certain microbes as secondary metabolites. Because of their value as bioactive natural products, there is significant interest in delineating the biosynthetic mechanisms of this unusual class of compounds. The primary difference between the biosynthetic origins of marine vs non-marine isocyanides is that the former are derived from terpenoids while terrestrial isocyanides are mostly derived from amino acids19. Most of the backbone of these terrestrial isocyanides (the xanthocillins, the cyclopentanes and others), as well as the nitrogen atoms in the isocyanide group, originate primarily from tyrosine 21–28. However, the origin of the carbon atom of the isocyanide group remained a mystery for some time. Puar et al 21 attempted to address this issue by synthesizing hazimicins from cultures of Micrococcus luteus that were fed 14C-labelled precursors; their findings indicate that the isocyanide carbon in hazimicins originates from methionine 21–23,29. Investigations into the source of the isocyanide carbon atoms of
Xanthocillin X monomethyl ether (XME) in glucose-starved cultures of fungus
Dichotomomyces cejpii via [U-13C] glucose feeding experiments revealed that the isocyanide carbon atoms were derived from glucose metabolism rather than pathways 4 involving tetrahydrofolate metabolism such as methionine, glycine, serine, TCA cycle, cyanides, cyanates or carbamoyl phosphate22,29.
Fig. 2: Biosynthesis of isocyanides and origin of the isocyano group; the carbon atom of the isocyano group comes from ribulose-5-phosphate, the nitrogen atom can originate from tryptophan (A) or tyrosine (B). Chemical structures constructed using ChemDraw (Perkin Elmer).
The specific precursor of the isocyanide carbon remained unknown still. Interestingly,
Penicillium notatum produces a di formamide compound, which appears in the culture before Xanthocillin and, which when added to the fungal culture, boosts the production of Xanthocillin X2,30. This suggests that formamides might be the precursors of isocyanides via dehydration reactions. This notion is debatable, as the discovery of the enzyme isocyanide hydratase (ICH) shows that formamides can the metabolic products of isocyanide, rather than precursors. ICH hydrates the isocyano groups in isocyanides to synthesize the corresponding formamides, which are then degraded to the corresponding amines and formates31–33. Therefore, formamide is one product of isocyanide degradation rather than a synthetic precursor, although not all biosynthetic routes to isocyanides are known. 5
Clardy et al 34 made significant headway into solving the mystery of the origin of the carbon atom of the isocyanide group when they developed a novel approach of cloning and heterologous expression of eDNA. eDNA is DNA from environmental samples (uncultured soil bacterium) that can be transformed into easily cultured lab bacteria, permitting the identification and characterization of antibiotics from
“antibacterially active eDNA clones using a high-throughput phenotypic screen”. They isolated an eDNA clone, CSLG18, which produces an indole-based isocyanide in E. Coli and identified its biosynthetic genes, isocyanide synthase (IsnA) and ɑ ketoglutarate- dependent oxygenase (IsnB) (Fig. 2). They observed that all the nitrogen atoms in this novel isocyanide are derived from the amino acid tryptophan. The source of the carbon
35 atom of the isocyanide was identified as the C2 atom of Ribulose-5-phosphate . The biosynthetic reaction proceeds via attachment of the carbonyl carbon of ribulose-5- phosphate to the ɑ amine of L-Tryptophan, catalyzed by isocyanide synthase (IsnA). The resulting intermediate is then resolved to the isocyanide product by hydroxylation and decarboxylative desaturation by the oxygenase (IsnB) (Fig. 2). 34–36. During this process, the carbonyl carbon of ribulose-5-phosphate undergoes a 2-electron oxidation to become the isocyanide carbon atom, catalyzed by IsnA and B acting co-operatively 36.
1.1.1.b Identification of IsnA/B homolog gene clusters: The identification of biosynthetic genes and a biosynthetic pathway of an isocyanide was an important discovery. This opened the door to a large number of isocyanide biosynthetic studies involving IsnA/B homologs when a BLAST search of sequenced bacterial genomes revealed several IsnA homolog operons 37. One class of isocyanide natural products whose biosynthesis was clarified by the discovery of IsnA/B is the hapalindoles. 6
Hapalindoles are terpenoid indole-containing bioactive alkaloids produced by cyanobacteria belonging to the order of Stigonemataceae (Fischerella sp. and
Hapalosiphon sp.); these compounds are divided into several classes, including the welwitindolelinones 38,39 and
Fig. 3: The PvcABCD operon in Pseudomonas aeruginosa synthesizes the isocyanide metabolite, Paerucumarin and its corresponding hydration product, Pseudoverdine from Tyrosine and Ribulose-5-phosphate. Paerucumarin signals the upregulation of biofilm development gene cluster cupB/C in pathogenic P. aeruginosa either by directly targeting the histidine kinases RocS1 and S2 or indirectly by activating an undetermined transcriptional activator “Y”. RocS1 and RocS2 signaling results in activation of cupC/B genes. Chemical structures constructed using ChemDraw (Perkin Elmer).
. ambiguines40,41(Fig. 6). (covered in more detail below). Investigations into the biosynthetic genes for these isocyanide hapalindoles revealed a 42Kbp ambiguine (amb) biosynthetic gene cluster in cyanobacterium Fischerella ambigua UTEX1903. This cluster encode the enzymes AmbI1-3, responsible for the isocyanide group assembly 39.
Interestingly, AmbI1-3 are close homologs of IsnA/B (46% identity), with AmbI1-2 sharing homology with IsnA and AmbI3 with IsnB. AmbI1-2 can functionally complement IsnA in vitro, converting L-tryptophan and ribulose-5-phosphate into the same intermediate as in Fig. 2. This intermediate is then resolved by AmbI3 into Z-3-(2- 7 isocyanovinyl)-1 H indole, instead of the isomeric E-3-(2-isocyanovinyl)-1 H indole synthesized when the intermediate is resolved by IsnB 36. This indicates that IsnB is specific to the E isomer while AmbI3 is specific to the S form (Fig. 2). Investigations into the biosynthetic genes for welwitindolinone-type isocyanides revealed a 36Kbp gene cluster in Hapalosiphon welwitschii UTEX B183042 containing the proteins WelI1,2,3 which are close homologs of AmbI1,2,3 from the amb operon. Whether WelI1,2,3 proteins can attach the nitrogen atom from L-tryptophan with the carbonyl carbon from ribulose-5-phosphate to generate a vinyl isocyanide, as done by AmbI1,2,3, remains untested.
Other homologs of IsnA/B include PvcA/B; IsnA has a 33% sequence identity with pvcA which is involved in C-N bond formation34,43. IsnB shows sequence similarity to non-heme Fe2+ ɑ-ketoglutarate-dependent oxygenases, including pvcB 44. The PvcA and B genes are a part of the Pvc gene cluster, which is a four gene (Pvc A, B, C, D) operon in Pseudomonas aeruginosa. P. aeruginosa is a human pathogen and its in vivo growth and pathogenesis are promoted by the synthesis of two siderophores, pyoverdine and pyochelin43. Initially the Pvc gene cluster was implicated in the synthesis of the pyoverdine chromophore, hence the name of the operon, but later this claim was refuted as the major product of this operon is a cyclized tyrosine derived isocyanide containing cumarin, paerucumarin (2-isocyano-6,7-dihydroxycoumarate) 44,45. The biosynthetic reaction was proposed by Brady and Clarke-Pearson45 (Fig. 3). The pvc genes were also shown to be involved in the synthesis of pseudoverdine, a pyoverdine-related fluorescent bicyclic compound. Paerucumarin is an unstable compound and can easily decompose into pseudoverdine by hydration (the isocyanide group is simply replaced by N-formyl 8 group in pseudoverdine) 46. Interestingly, the Pvc gene cluster is negatively regulated by the presence of iron. Under conditions of iron limitation, the Pvc operon is upregulated to either synthesize the siderophore pyoverdine in certain strains of Pseudomonas, or to synthesize paerucumarin and pseudoverdine, which are potentially siderophores.
Moreover, the Pvc gene cluster in P. aeruginosa is positively regulated by the LysR family transcription factor PtxR 45, which also upregulates the expression of the toxA gene encoding exotoxin A under conditions of iron limitation47. In addition, PtxR regulates the expression of other virulence factor genes that promote quorum sensing in
P. aeruginosa48. However, the role of the Pvc gene cluster in promoting P. aeruginosa pathogenicity remains unexplored. Moreover, PvcA and B homologs are found in a large number of human, insect and plant pathogenic microbes, such as, Legionella pneumophila, Vibrio cholerae, Burkholderia mallei, Photorhabdus luminescens,
Xenorhabdus nematiphila, Erwinia carotovora, Bdellovibrio bacteriovirus, etc. 45.
Legionella pneumophila, the causative agent of Legionnaire’s disease, is an infectious bacterium that possesses a five gene biosynthetic gene cluster (BGC) containing homologs of isnA and B named lpg0174 and lpg0175 49. Moreover, this gene cluster is upregulated during the growth of the bacteria in human host macrophages, which suggests its potential role in virulence, possibly as siderophores to promote iron uptake by the pathogen. The BGC operon encodes four novel N-acyl-L-histidines
(primarily by the lpg0178 gene), consistent with the observation 50 that L-histidine biosynthesis in upregulated during the growth of the pathogen within human macrophages. 9
E. coli cells expressing the Legionella BGC operon (lpg0173-177) were capable of synthesizing the isocyanide rhabduscin (Fig. 2) 49 51, although rhabducin was not detected in wild type Legionella cells. Possible explanations for this are that rhabducin is unstable and rapidly converted to some other untraceable metabolite or that other experimental conditions are required for expression of the BGC biosynthetic operon.
Rhabduscin is also produced by the insect pathogens Xenorhabdus nematophila and
Photorhabdus luminescens. The Xenorhabdus nematophila ATCC19061 genome contains a rhabduscin biosynthetic gene cluster comprising the IsnA and B homologs
XNC1_1221 and XNC1_1222 as well as a glycosyltransferase gene (XNC1_1223) 51, similar to a biosynthetic gene cluster in Photorhabdus luminescens TT01 51.
Heterologous expression of the rhabduscin biosynthetic transgenes in E. coli resulted in the synthesis of the minor isocyanide functionalized glycoside, byelyankacin, and rhabduscin, although the latter could not be detected in most cases owing to sensitivity of the isocyanide group to hydrolysis to yield formamide derivatives. It would be interesting to know if hydrolysis of the isocyanide group in rhabduscin in E. coli is controlled by a
Pseudomonas ICH homolog enzyme.
Byelyankacin, or E-4-(2-isocyanivinyl) phenyl ɑ-L-rhamnopyranoside, was first isolated from Enterobacter sp, and was shown to be an inhibitor of phenoloxidase-type enzyme tyrosinase and melanin synthesis in humans51,52. This, along with the observation that isocyanides are produced by a large number of pathogens, has directed attention towards the potential role of Rhabduscin, and other isocyanide natural products, in the suppression of host innate immunity. 10
Fig. 4. Role of (left) ScoB-E (Actinomycetes sp) and (right) MmaB-E (Mycobacterium marinum) in INLP1-2 biosynthesis.
.
In addition to the discussed mechanism of biosynthesis of isocyanides involving IsnA and B and their homologs, another mechanism was proposed involving a biosynthetic gene cluster in Mycobacterium. Mycobacterium tuberculosis, the causative agent for tuberculosis, contains a biosynthetic operon, Rv0096-0101, encoding five enzymes conserved across pathogenic mycobacterium as well as Streptomyces, Nocardia,
Rhodococcus etc. These enzymes are an iron(II) and α-ketoglutarate (α-KG)- dependent oxidase, fatty acyl-CoA thioesterase, acyl-acyl carrier protein ligase (AAL), acyl carrier protein (ACP) and non ribosomal peptide synthase. Co-expression of these genes from
Mycobacterium marinum (mma-E) and Streptomyces coeruleorubidus (scoA-E), in E. coli resulted in the synthesis of two isocyanide-functionalized lipopeptides (INLP 1 and
2). INLP1 and INLP2 are structurally similar to two known isocyanide antibiotics
(SF2768 and SF2369) from Actinomycetes sp 53. Investigation into the biosynthetic 11 mechanism revealed a multistep process (Fig. 4), starting with the adenylation of crotonic acid by ScoC, which is then transferred to an acyl carrier protein (ScoB); a lyase (ScoD) attaches a glycine to the β position of the ɑ, β unsaturated fatty acyl-ACP which then undergoes oxidation and dephosphorylation by the oxidase (ScoE) to form the isocyanide group. A similar biosynthetic mechanism exists in M. marinum with MmaC that favors longer chained fatty acid substrates, in contrast with ScoC53(Fig. 4).
The previously mentioned di-isocyanide antibiotic SF2768, is produced by the
Actinomycetes Streptomyces thioluteus, which contains an isocyanide biosynthetic operon (sfa) encoding a dioxygenase (SfaA), an AMP ligase (SfaB), a peptidyl carrier protein (SfaC), a nonribosomal protein synthetase (SfaD) and a hydroxylase (SfaE) 54
(Fig. 5). Interestingly, production of this di-isocyanide is often accompanied by mixed isocyanide-formamides and even di-formamides; this led to the discovery of another gene in the sfa operon, SfaF that encodes an isocyanide hydratase. Isocyanide hydratase adds water to the isocyano groups to yield formamides (SfaF has a 32% sequence identity to
InhA, encoding the first characterized isocyanide hydratase from Pseudomonas putida.
32,55.
The identification of so many isocyanide synthase (IsnA/B) homologs in sequenced bacterial genomes suggests the possibility that several more isocyanide biosynthetic gene clusters reside in the genomes of uncultured bacteria. A total of 12
IsnA containing biosynthetic gene clusters were identified and isolated from approximately 400,000 screened eDNA cosmid clones. Some of these operons contained simply the IsnA and B domains, while others were more complex and contained additional genes encoding biosynthetic enzymes 37. The gene clusters were amplified and 12 expressed in E. coIi or Pseudomonas aeruginosa to identify the clone-specific metabolites, 4 of which were shown to contain isocyanide groups 37. Importantly, one of these four isocyanides is Paerucumarin, a previously described isocyanide natural product
(Fig. 3).
The first isocyanide biosynthetic pathway in a fungus, Aspergillus fumigatus, was identified very recently56. The genome of Aspergillus fumigatus AF293 contains four isocyanide synthases (ICS), namely, CrmA, IcsA, IcsB and XanB, spread between three biosynthetic gene clusters (ICS3G, ICS4G and ICS5G). Under conditions of copper limitation, the ICS5G gene cluster containing the isocyanide synthase XanB produces four Xanthocillin derived isocyanides in A. fumigatus: a diformamide, a fumiformamide,
BU 4704, and Xanthocillin monomethyl ether (XME)56 (Fig. 6). Inspection of the
Xanthocillin biosynthetic pathway in A. fumigatus reveled a multistep process beginning with the conversion of L-tyrosine to an intermediate by XanB, similar to isocyanide synthase IsnA/PvcA in bacteria; this intermediate then undergoes an oxidative dimerization to Xanthocillin catalyzed by a cytochrome P450, XanG; Formyl and methyl derivatives of Xanthocillin are synthesized by XanA (homologue of InhA from
Pseudomonas sp) and the methyltransferase XanE respectively56 (Fig. 2.B). An analogous biosynthetic pathway has been observed in yeast Saccharomyces cerevisiae, involving the sporulation specific genes, Dit1 and Dit2. Dit1, which is homologous to
XanB, converts tyrosine into an isocyanide-functionalized, tyrosine-derived precursor and ultimately to N-formyl tyrosine; this is then converted to N,N-bisformyl dityrosine by Dit2 which is a member of the cytochrome P450 family and a XanG homolog56–58
(Fig. 2.B). It is important to note here that the existence of the isocyanide formed in the 13 first step of the biosynthetic reaction catalyzed by Dit1 has not been observed in yeast, although it is entirely possible that the isocyanide group is rapidly hydrated to formyl group by a hydratase in yeast. This provides mechanistic rationale for the observation that isocyanide synthases and isocyanide hydratases often exist in pairs and the synthesis of the latter is often induced by the presence of isocyanide32. A better understanding of isocyanide biosynthesis and metabolism may elucidate the ways in which soil-dwelling microbes balance defense and stress response in the soil microbiome.
1.1.1.c Biological targets of terrestrial isocyanides: The pvcA-D gene cluster in
Pseudomonas aeruginosa encodes the isocyanide natural product paerucumarin, as mentioned previously. Interestingly, mutations in the pvc gene operon in P. aeruginosa
MPAO1 resulted in downregulation of the expression of another gene cluster, the cup operon (cupB, cupC). Cup proteins are responsible for attachment to abiotic surfaces and biofilm development in various infection sites. This indicates that the pvc gene cluster controls biofilm formation in P. aeruginosa by regulating the cup gene operon using paerucumarin as a signaling molecule59. The transcriptional factor Ptxr upregulates the pvc gene cluster to synthesize paerucumarin, which in turn activates either the CupB/C two component regulatory system or the roc system. This occurs with the help of histidine sensor kinases, RocS1 and RocS2, which either directly activate RocA1 and an unknown effector “X” or indirectly via, an unknown transcriptional activator “Y”; this in turn upregulates cupC1-3 and cupB1-6 expressions respectively59,60 (Fig. 3). 14
Fig. 4: Isocyanide SF2768 is synthesized by the SfaABCDE operon in S. thioluteus. This isocyanide is a chalkophore; it is transported outside the cell membrane by Orf12 where in, it chelates copper (I) and transports it into the cell via ABC transporters (Orf 9-21). Inside the cell, the chalkophore bound copper is released and SF2768 is regenerated for another cycle of copper chelation.
There is an intriguing possible connection between isocyanide natural products and biofilm formation mediated by iron availability. Biofilm development in several pathogenic bacteria, such as L. pneumophila 61, P. aeruginosa 62, S. aureus 63,64, etc, is induced by low levels of intracellular iron. Under conditions of iron limitation, P. aeruginosa synthesizes two endogenous siderophores, pyoverdine and pyochelin, which are responsible for iron acquisition from the host 65 and subsequent biofilm development around infection sites66–68. As mentioned previously, under conditions of iron limitation the Pvc gene cluster directs the synthesis of the siderophores pyoverdine and pyochelin as well as the isocyanide paerucumarin and its formyl form pseudoverdine 45. It would be interesting to determine if paerucumarin and pseudoverdine can chelate iron, possibly pointing to an interaction between these isocyanide and isocyanide-derived natural 15 products and the expression of genes critical for biofilm development in P. aeruginosa62,69. If validated, isocyanide-induced biofilm development would be a novel mechanism for biofilm formation in P. aeruginosa and other pathogenic strains possessing the pvcA-D operon homologs. Moreover, Ferric uptake regulator (Fur) is an iron dependent transcription factor that regulates the expression of genes critical for biofilm development, and the major intracellular iron regulator in P. aeruginosa62,69.As such, it would be interesting to analyze the interplay between the cup genes (CupB1-6 and CupC1-3) and the fur responsive genes in promoting pathogenesis and biofilm synthesis.
The notion that isocyanides may act as siderophores is supported by their intrinsic ability to form complexes with a variety of transition metals70,71, inherent affinity towards hemoproteins72,73, and also by a large body of correlative experimental evidence suggesting that pathogenic bacteria synthesize isocyanides to chelate metals and promote survival in hosts. As mentioned previously, Mycobacterium tuberculosis, the causative agent for tuberculosis, contains the Rv0096-0101 biosynthetic operon that encodes two isocyanide-functionalized lipopeptides,INLP1 and 2; these two show structural homology to isocyanide antibiotics SF2768 and SF2369 from Actinomecetes sp53. This conserved biosynthetic operon (Rv0096-0101) appears critical for survival and pathogenesis of M. tuberculosis in macrophages. While the mechanism of this remains undiscovered, there is the possibility that the Rv0096-0101 encoded genes facilitate metal uptake, transport, and homeostasis in the pathogenic bacteria.
Additional evidence for an isocyanide-transition metal connection is provided by
CtpA and CtpB: two heavy metal-translocating ATPases located near the isocyanide 16 biosynthetic gene cluster in the genome of M. tuberculosis. The expression of CtpB is co- upregulated with the isocyanide biosynthetic gene cluster (Rv0096-0101) under conditions of zinc limitation 53,74,75. Deletion mutations in this conserved isocyanide biosynthetic gene cluster (mmaA-E) in the pathogenic mycobacterium M. marinum resulted in a significant drop in intracellular zinc accumulation, consistent with the observation that this isocyanide biosynthetic gene operon is important for metal uptake and transport in pathogenic mycobacterium53. However, there is currently no direct evidence of metal chelation by the encoded isocyanide lipopeptides and a detailed mechanism of metal uptake and the resulting interactions between the isocyanides and the components responsible for regulating metal homeostasis in these pathogens is unknown.
There is a more direct connection to metal binding in the isocyanide antibiotic
SF2768 produced by Streptomyces thioluteus. SF2768 is a chalkophore, that is, a molecule that binds to and facilitates acquisition of extracellular copper. To acquire extracellular copper, SF2768 (Fig. 5) is transported across the cell membrane by Orf12, a member of the major facilitator superfamily (MFS) exporters. SF2768 chelates and converts environmental copper (Cu2+) to Cu+ to form an SF2768-Cu+ complex, which is then transported inside the cell by an ABC transporter encoded by genes Orf9-21 in the isocyanide biosynthetic (sfa) gene cluster. Once inside the cell, the chelated Cu+ is released binds to several Cu-dependent enzymes critical for growth and development as well as secondary metabolite synthesis (by tyrosinase-like copper containing monooxygenases) in Streptomyces species 54,76–78.The SF2768 is regenerated and ready for another cycle of copper acquisition54 (Fig. 5). SF2768 derivatives are also copper chelators; the isocyanides have the highest copper chelating activities, followed by the 17 partially hydrated metabolites. The diformamides have no detectable copper chelating activity, indicating that the presence of isocyano group(s) is directly responsible for copper binding and transport 55.
Unsurprisingly, SF2768 and its derivatives are not the only instances of isocyanides potentially acting as chalkophores. Human pathogen Aspergillus fumigatus
AF293 genome contains four isocyanide synthases (ICs), CrmA, IcsA, IcsB and XanB, spread across three biosynthetic gene clusters (ICS3G, ICS4G and ICS5G), as mentioned previously 56. The biosynthetic gene cluster ICS3G, consists of four genes (crmA, B, C and D), referred to as the crm cluster, which are expressed only in copper limited conditions and their expression patterns are strikingly similar to that of high affinity copper transporter ctrC. The expression of crmA-D is not upregulated in iron deplete conditions, thus indicating their specific transcriptional response to copper. MacA, AceA and CufA are three copper binding transcription factors present in most pathogenic fungi, including Aspergillus sp, involved in copper uptake during copper depletion, detoxification of copper at toxic levels and both, respectively56,79. MacA induces the expression of the crm cluster under copper depleted and replete conditions56 while AceA and CufA has no effect. Interestingly, out of the four crm genes, crmC and crmD encode copper and iron transporters respectively, but the mechanism of copper or iron uptake and transport by crmD and crmC encoded transporters remain unclear.
Promoting virulence of a pathogen in a host requires survival of the pathogen inside the host. An important determinant of pathogenicity is to overcome the host’s natural immune defense. 80 One commonly used mechanism for subverting the host defense is disruption of host redox homeostasis79,81. Isocyanides have been shown to 18 promote pathogenicity by inhibition of copper-containing enzymes. Copper plays an essential role in the host’s innate immunity, either by serving as cofactor to physiologically important enzymes, promoting differentiation, maturation and proliferation of leukocytes, or by ensuring proper neutrophil and macrophage functions against pathogens82–86. Therefore, isocyanides chalkophores can inactivate the host’s innate immune defense machinery by disruptive copper sites in these proteins.
An example of an isocyanide that inactivates copper-dependent enzymes involved in mediating proper host immune functioning is rhabduscin. Rhabduscin is synthesized by insect pathogens Xenorhabdus nematophila and Photorhabdus luminescens (Fig. 2.B) and can inhibit the copper-dependent enzymes phenoloxidase and tyrosinase at nanomolar concentrations 51. These metalloenzymes contribute to the immune defense of an insect host by synthesizing and depositing melanin, which insects use as protective layer to seal out pathogens87,88. Therefore, rhabduscin promotes pathogenesis both by disrupting the host’s ability to synthesize melanin and through its antimicrobial properties that kill competitors and enhance pathogen survival in host insect cadavers.
Other isocyanides also inhibit tyrosinase. For example, melanocin A (Fig. 6) an isocyanide antibiotic isolated from Eupenicillium shearii, inhibits mushroom tyrosinase and subsequently melanin biosynthesis (IC50 of 9nM). Structurally related compounds lacking the isocyanide functional group, namely, Melanocin B and C, failed to inhibit tyrosinase, thus indicating that the isocyanide group is essential for inhibition89. Several cyclopentyl-type isocyanides from Trichoderma harzianum MR304, were all inhibitors of melanogenesis in Streptomyces bikinienesis and B16 melanoma cells; they inhibited mushroom tyrosinase, and consequently melanogenesis, with IC50 values of 0.25 µg/ml 19 and 0.05µg/ml respectively 90. The precise mechanism by which isocyanides inhibit tyrosinase is unclear, but competitive binding of the isocyano group of the inhibitor to the copper in the active site of tyrosinase has been speculatively proposed 91. It is not known if there are isocyanides that inhibit metalloenzymes involved in the human immune system or are implicated in protection against other human diseases, which is an avenue for future research.
1.1.1.d Types of terrestrial isocyanides (Fig. 6): i. Xanthocillin-class isocyanides: Xanthocillin was isolated from Penicillium notatum by Rothe, as a complex mixture of xanthocillins X, Y1, Y2 and Z. The members of this group are all structurally related to xanthocillin (Fig. 6), with variations in the groups on their aromatic rings 2. Interestingly, studies show that xanthocillin has also been produced outside the planet Earth, in space; Penicillium expansum 2-7 and Penicillium chrysogenum, resident strains of the orbital complex, Mir, which became the dominant strains at the end of a long term space flight, produced secondary metabolites, xanthocillin X and questiomycin A92,93. Aspergillus is another important source of xanthocillin and its derivatives 94,95,96 such as xanthocillin X dimethylether (XDE) and xanthocillin X monomethylether (XME) (Fig.6). XME has also been isolated from
Dichotomomyces cejpii, where it acts as an inhibitor of prostaglandin biosynthesis and platelet aggregation 94,97. Other important xanthocillin-derived isocyanides include xanthoascin, isolated from Aspergillus candidus 2,94,98 and emerin, a naturally existing di- cyano analogue of xanthocillin2, isolated from the mycelium of Aspergillus nidulans 94,99
(Fig. 6). BU 4704 is another xanthocillin-type isocyanide which was isolated from
Aspergillus sp by Tsunakawa et al 100. Aspergillus fumigatus co cultured with 20
Streptomyces peucetius resulted in the synthesis of a formyl xanthocillin analogue, known as fumiformamide 101.
Biological activity of Xanthocillin class isocyanides:
Fig. 5: TOP: Xanthocillin type isocyanides; BOTTOM: Cyclopentyl type isocyanides. Together, xanthocillins and cyclopentyls cover the largest percentage of non-marine isocyanide natural products. Structures constructed using ChemDraw (PerkinElmer).
Xanthocillins as antibacterial and antiviral agents: Isocyanides are natural products produced by certain microbes in secondary metabolism and almost all of them are potent antibiotic agents. The Xanthocillin complex (X, Y1, Y2, Z) exhibits a wide range of activity against both gram positive and negative bacteria as well as fungi and yeasts2.
This broad spectrum antimicrobial property of Xanthocillin may be the reason that the producer fungal strain, Penicillium, was the dominant surviving fungus on the Mir space station after long term flight92,93.Three di-hydrogenated forms of Xanthocillin, namely
NK372135 A, B and C, isolated from the fungus Neosartoria fischeri var. glabra
IFO9857, strongly inhibits the growth of Candida albicans in vitro with IC50 values of 21
2.12, 0.53 and 0.27 µg/ml respectively 102, thus providing potential therapies against candidiasis.
The xanthocillin mono methylester (XME), xanthocillin dimethylester (XDE) and methoxy xanthocillin dimethylether derivated isolated from Apergillus sp. exhibit anti- viral activity. These xanthocillin derivatives are active against Newcastle disease virus
(NDV), the causative agent for Newcastle disease, a contagious bird disease that is easily transmissible to humans. Xanthocillin esters are effective against herpes simplex virus and Bacillus subtilis 103. They are, however, not effective against the plant pathogen tobacco mosaic virus (TMV) 104. This suggests that xanthocillin-derived antibiotics may be generally less effective against plant viruses, potentially owing to impaired diffusion of these antibiotics across the protective plant leaf cuticle layer104.
More studies are required to confirm this hypothesis.
The mechanism of action of XME against NDV in cultured cells involves the reversible inhibition of the synthesis of hemagglutinnin in NDV-infected cells 95 at 15
µg/ml concentration. XME is one of the most effective antiviral inhibitors of protein synthesis, effectual at low concentrations and with low cytotoxicity 95. Identification of
XME’s physiological targets, inhibitory mechanism against hemagglutinin synthesis, and role in prevention of hemagglutination requires further study.
A new type of xanthocillin-like antibiotic was isolated from Leptosphaeria sp. L-
179 in 1990 and named MK4588 or Leptocillin. MK4588 shows strong activity against several infectious bacterial species (Klebsiealla, Proteus mirabilis, Providencia rettgeri,
Morganella morganii kono) at concentrations ranging from 0.78-12.5µg/ml, but no activity against E. coli at concentrations as high as 100µg/ml 105. Two other xanthocillin- 22 derived compounds, darlucins A and B (Fig. 6), were isolated from Darluca filum. They exhibit antibacterial (MIC: 2.5-20µg/ml), antifungal (MIC: 2.5-50µg/ml), as well as low cytotoxic effects against various cell lines, and no phytotoxic effects 106.
Xanthocillins inhibit prostaglandin biosynthesis: Prostaglandins are a family of fatty acid-derived molecules that are powerful vasodilators and are primarily involved with causing fever, pain, and inflammation. In 1982, Vane 107 won the Nobel prize in medicine for his discovery of the mechanism of action of non-steroidal anti-inflammatory drugs or
NSAIDs in the inhibition of prostaglandin biosynthesis, The classical NSAID Aspirin
(acetyl salicylic acid) inhibits the enzyme cyclooxygenase (COX), thereby preventing the conversion of the fatty acid (arachidonic acid) to prostaglandins, thus providing relief from pain and inflammation108. Around the same time, Kitahara et al 97 observed that xanthocillin monomethylether (XME), isolated from cultures of Dichotomomyces cejpii, reduced prostaglandin (specifically prostaglandin H2) biosynthesis from Arachidonic acid by 50%. The mechanism of inhibition of PGH2 biosynthesis by XME is unknown.
CSBP is a protein kinase, involved in the biosynthesis of inflammatory cytokines
IL-1 and TNF as a response to a large number of inflammatory signals. There are several classes of imidazole CSBP ligands which are capable of binding to CSBP and inhibiting
LPS stimulated synthesis of inflammatory cytokines. They also show variable extent of inhibition of arachidonic acid metabolism by inhibiting the enzymes prostaglandin H synthase 1 (PGH1) or COX1 and Arachidonate 5-lipoxygenase (5-LO) 109. Boehm et al synthesized a new class of CSBP ligand, 1substituted-4-aryl-5-piridinylimidazoles from isocyanides (tolyl sulfide isocyanide) utilizing van Leusens’ methodology 110 and observed that it inhibits 5-LO and Cycloxygenase activity with low potency109. 23
Nevertheless, this warrants investigations into the potential inhibitory effect of isocyanides on cyclooxygenase activity and their possible utility as anti-inflammatory drugs and analgesics.
Xanthocillins are thrombopoietin receptor agonists: Thrombopoietin (TPO) is a cytokine that is responsible for the regulation of megakaryocyte and platelet production via activation of its receptor, MPL or TPO-R 111,112. Recombinant human TPO (rh-TPO) is currently used for the treatment of chemotherapy-induced thrombocytopenia, a potentially fatal condition characterized by low platelet count 113,114. However, some serious limitations with the use of rh-TPO include development of neutralizing antibodies against TPO and limitation with oral administration (rh-TPO- in unstable in the intestine)
115,116. Sakai et al, reported that four xanthocillin derivatives, including XDE as the first natural TPO mimetics, These exhibit TPO receptor agonist activity115,117 and promote the proliferation of TPO-sensitive human leukemia cell line (UT-7/TPO) in a dose dependent manner. Yamaguchi et al studied the contribution of the chemical structure of xanthocillin derivatives towards their TPO-R agonist activity. They observed that TPO-R agonist activity of these isocyanides are enhanced by alternations in the olefin geometry and by hydrophobic substitutions on benzene rings 115 but high concentrations of these compounds inhibit cell proliferation, potentially due to the cytotoxicity of vinyl isocyanide groups required for their bioactivity. More dose optimization and structure- activity studies of these TPO mimetics are need in order to diminish their toxic effects and to generate potent replacements of rh-TPO in the treatment of thrombocytopenia.
Complications from the hepato- and cardio-toxicity of Xanthocillins: Townsend et al classified xanthocillin as “too hepatotoxic for use” 118. Xanthoascin (Fig.5). isolated from 24
Aspergillus candidus, has been shown to exhibit hepato- and cardio-toxicity and to induce teratogeneicity in mice 119,120. It also causes severe hepatic injury, jaundice, and focal or confluent necrosis of hepatocytes in mice121,122 , followed by myocardial degeneration. In addition, xanthoascin generates unique lesions in the liver and the heart, which are caused by vacuolation of the lung alveolar and myocardial interstitial cell nuclei 122. Since Aspergillus candidus is an extremely common soil fungus known to contaminate food, xanthoascin poses a serious risk of food borne liver and heart diseases.
While xanthoascin poses a potentially serious human health risk, there might be other bioactive isocyanide products, such as the darlucins, which exhibit low cytotoxicity against various cell lines, and no phytotoxic effects 106; these isocyanides are attractive targets for therapeutic interventions and drug design. Several isocyanides, such as, brasilidine A, from Actinomycete Nocardia brasiliensis 123, exhibits significant cytotoxicity against various tumor and multidrug resistance cell lines124 and can be used as anti-cancer agents.
Xanthocillins as anti-malarials and inhibitors of melanin biosynthesis: The xanthocillin-type alkaloid, cordyformamide, was isolated from the fungus Cordyceps brunnearubra and is toxic to the malarial parasite Plasmodium falciparum but has low toxicity to human breast cancer cells. It is not cytotoxic towards other cancer cells, such as small cell lung cancer, epidermoid carcinoma, etc 125. Cordyformamide has a structure similar to that of melanocin A, a melanin biosynthesis inhibitor (IC50 of 9nM) isolated from the fungus Eupenicillium shearii 89. Cordyformamide possesses two formamide groups instead of the isocyanide group found in xanthocillin Y2, while melanocin A possesses one formamide and one isocyanide group (Fig.5). 125. Interestingly, melanocin 25
B and C, which are structurally related to Melanocin A but do not possess isocyanide groups, failed to inhibit tyrosinase. Because tyrosinase is involed in the biosynthesis of melanin, the isocyanide moiety in the compounds appears to be critical for their inhibitory effect on melanin production89.
Xanthocillins as anti-tumor agents: Xanthocillin dimethyl ether (XDE) inhibits the proliferation of liver HepG2, breast cancer MCF-7, and human oral carcinoma KB cells
126 with IC50 values of 0.18, 0.38 and 0.44 µg/ml respectively . Investigation into the mechanism of action of XDE against tumor cells revealed activation of pro-apoptotic pathways and increased levels of reactive oxygen species (ROS) 126. It is possible that
XDE targets ROS-regulated apoptotic pathways in cancer cells. A xanthocillin-derived compound isolated from the marine organism Penicillium commune, SD118-xanthocillin, inhibits the growth of a large number of cancer cell lines in a dose dependent manner. Its mechanism involved inducing autophagy by downregulating the MEK/ERK pathway along with upregulating PI3K/Beclin 1 signaling 127. ii Cyclopentyl-type isocyanides: Trichoderma, a common soil fungus, is the primary source of all cyclopentyl-type isocyanides. The first isocyanide of this type to be reported was dermadin (previously known as U-21963). It was isolated from Trichoderma viride in 1966 128,129, almost 10 years after the discovery of Xanthocillin in the fungus
Penicillium notatum. The members of this group of isocyanides vary primarily in their oxidation levels and degrees of hydration 2 (Fig.5). Dermadin and another related isocyanide, Trichoviridine, were also isolated from Trichoderma koningii 130 128.
Trichoviridine has also been isolated from various other sources and called different names, including compound 142B from Tricoderma sp, and isonitrin B from strain 26
OMRL 3200 (Trichoderma hamatum) 2,128,131. An isomer of isonitrin B has been isolated from Trichoderma harzianum l2. Isonitrins A and D were isolated from Trichoderma harzianum and Trichoderma hamatum respectively 2,132. Two new cyclopentyl-type isocyanides were isolated from Trichoderma hamatum133 , a spirolactone and a methyl ester2 (Fig.5). Several volatile isocyanides have been isolated from Trichoderma koningee, named homothallin 1, homothallin 2, and their related amine and N-formyl derivatives 134 2,135. Homothallin 2 has also been isolated from a mutant strain of
Trichoderma harzianum (the wildtype strain was not a producer of homothallin 2) 136.
Biological activity of Cyclopentyl-type isocyanides:
Antibacterial and antifungal properties of cyclopentyls: Cyclopentyl-type isocyanides are antibacterial (effective against both Gram negative and positive) and antifungal agents, similar to the previously discussed xanthocillins2,132. Trichoderma, the primary source of these cyclopentyl-type isocyanides, is widely used as a biocontrol agent; it inhibits the growth of other organisms in its vicinity. It is possible that the biochemical mechanism of this competitive advantage might include the biosynthesis of toxic secondary metabolites including isocyanides 135,137.
Cyclopentyl isocyanides contribute to ill thrift disease and alteration of rumen bacterial metabolism: “Ill thrift” (failure to thrive or FTT) is a common disease in ruminating animals 138, and is characterized by impaired growth, muscular dystrophy, anemia, and ketosis of the affected animal. Ovine ill thrift is contracted when sheep feed on pastures contaminated with Trichoderma. Cyclopentyl-type isocyanides secreted by
Trichoderma sp, such as trichoviridin, dermadin, and 3-(3-isocyanocyclopent-2- enylidene)-propionic acid (Fig.5) from Trichoderma hamatum 139,140 cause ill thrift 27 disease. The primary target of these Trichoderma metabolites is the bacteria that digest the cellulose in ruminating animals. The cyclopentyl isocyanides deplete beneficial rumen bacteria in the affected animal, causing disease 141. The inhibitory mechanism of these toxic Trichoderma isocyanide metabolites on beneficial rumen bacteria has been studied extensively. 3-(3 isocyanocyclopent-2-enylidene)-propionic acid induces carbohydrate fermentation changes in the rumen that are related to ill thrift disease 142,143 by increasing the production of acetate from carbohydrates for most of the 11 tested strains of rumen bacteria. However, some strains showed a net decrease of acetate production relative to the synthesis of other acidic fermentation products. 3-(3 isocyanocyclopent-2-enylidene)-propionic acid also causes nutritional deprivation of the animal 142.
Digestion of the cellulose in timothy hay, a type of grass was inhibited by 2-5
µg/ml of 3-(3-isocyanocyclopent-2-enylidene)-propionic acid in a mixed culture of rumen bacteria. 12 µg/ml of 3-(3-isocyanocyclopent-2-enylidene)-propionic acid inhibited the fermentation activity of rumen fluid, measured by its dehydrogenase activity
(DHA); DHA of rumen fluid decreases by 50% upon a 30-minute incubation with this metabolite, thus indicating that the primary inhibitory step is extremely rapid 144.
Interestingly, this inhibitory effect of the isocyanide on rumen fluid fermentation and cellulose digestion by rumen bacteria is significantly reduced by the introduction of equimolar amounts of Nickel (Ni2+) and Cobalt (Co2+) respectively, into the system.
Palladium and chromium ions are also capable of reducing the inhibition. Given the known affincity of isocyanides for metals, these metal ions may polymerize144 the isocyanide, thereby reducing its concentration and anti-bacterial effects. 28
One of the causes of ill thrift is cobalt deficiency 145. Cobalt is converted to
Vitamin B12 (cobalamin) by the rumen bacterial flora of ruminating animals 146. A deficiency in the production of vitamin B12 resulting from lack of cobalt causes impaired propionic acid metabolism in the liver, hepatic dysfunction, suppressed appetite, and impaired growth, ultimately resulting in ill thrift 147,148. This finding is consistent with other results that point to transition metal biochemistry as an important axis of isocyanide natural product activity.
Cyclopentyl isocyanides inhibit pleiotropic resistance (PDR) in bacteria and possibly multidrug resistance (MDR) in humans: One of the major persistent issues with the use of anti-cancer drugs is a phenomenon called multidrug resistance (MDR). In MDR, the ATP binding cassette (ABC) transporter proteins expel a wide variety of anti-cancer drugs from the cells. Because MDR reduces chemotherapy efficacy, developing inhibitors of these drug efflux pumps is a high priority 149. Pleotropic drug resistant protein 5 (Pdr5p) is an ABC transporter that acts as a drug efflux pump and is implicated in pleiotropic resistance (PDR) in Saccharomyces cerevisiae. This phenomenon is functionally similar to multidrug resistance (MDR) in mammals caused by the ABC transporter protein P-glycoprotein 150,151. The cyclopentyl-type isocyanide 3-(3- isocyanocyclopent-2-enylidene)-propionic acid was isolated from cultures of
Trichoderma sp. P24-3 while screening for inhibitors of MDR in tumors. In Pdr5p overexpressing cells, this isocyanide is capable of inhibiting Pdr5p-mediated efflux of cerulenin , a fatty acid synthase inhibitor 152. 3-(3-isocyanocyclopent-2-enylidene)- propionic acid is also capable of inhibiting efflux by the Pdr5p homologs Cdr1p and
Cdr2p in the pathogenic yeast, Candida albicans. 151 The mechanism of inhibition of 29
Pdr5p by the isocyanide is unknown, although it does not appear to be due to the downregulation of PDR5 gene expression or Pdr5p protein levels. Therefore, isocyanide inhibition must specifically targeting Pdr5p function 151. There is also a need to determine the effect of 3-(3-isocyanocyclopent-2-enylidene)-propionic acid on mammalian drug efflux pumps, and to determine the role of this cyclopentyl-type isocyanide in overcoming multidrug resistance in humans as a viable cancer treatment strategy.
Cyclopentyl isocyanides inhibit tyrosinase and melanin biosynthesis: Tyrosinase is a copper-dependent enzyme found in both prokaryotes and eukaryotes that is responsible for the biosynthesis of melanin 91. During the late 1990s and early 2000s, many research groups focused on screening microbes for the production of secondary metabolites which inhibit the process of melanin synthesis (melanogenesis) by directly inhibiting tyrosinase
153. Several tyrosinase inhibitors were isolated 90 from Trichoderma harzianum MR304, two of which were novel. All of the identified inhibitors were cyclopentyl isocyanide compounds and inhibited melanogenesis in Streptomyces bikinienesis and B16 melanoma cells with submicromolar IC50 values. The two newly discovered isocyanides were named
MR566A or 1-(3-chloro-1,2 dihydroxy-4-isocyano-4-cyclopenten-1-yl) ethanol and
MR566B or 1-(1,2,3—tri-hydroxy-3-isocyano—4-cyclopenten-1-yl) ethanol. The other isolated isocyanides included 1-(1,4,5-trihydroxy-3-isocyanocyclopenten-2- enyl)ethanol, 2-hydroxy-4-isocyano-α-methyl-6-oxabiocyclo[3.1.0]hex-3-ene-
3methanol, 4-hydroxy-8-isocyano-1-oxaspiro[4.4]cyclonon-8-en-2-one, MR304A, methyl-3-(1,5-dihydroxy-3-isocyanocyclopent-3-enyl)prop-2-enoate90,90,154. The mechanism of inhibition of tyrosinase by these isocyanides is unknown, although the copper in the active site of the tyrosinase enzyme is a likely target. The inhibition of 30 tyrosinase by several isocyanides, including the homothallin 2 structural analog isolated from marine Trichoderma viride H1-7 91,155, suggests a shared mechanism.
Possible involvement of cyclopentyl isocyanides in fungal reproduction:
Phytophthora cinnamomi Rands is a soil-borne fungus and a plant pathogen responsible for the widespread root rot disease in plants. This fungus is heterothallic with compatible mating strains A1 and A2 156. Brasier 157,158 observed that certain volatile metabolites from Trichoderma viride were able to stimulate oospore production by the A2 mating type and not the A1 mating type of Phytophthora cinnamomi Rands following a homothallic pattern. Pratt et al 134 observed the same oospore induction in the A2 mating strain of Phytophthora cinnamomi Rands by metabolites from Trichoderma koningii.
These volatile metabolites from Trichoderma koningii were identified as the cyclopentyl isocyanides homothallins 1 and 2 2. The mechanism of induction of homothallic oospores in Phytophthora cinnamomic A2 mating type by homothallins remains unclear. iii Assorted other terrestrial isocyanides: Non-marine natural isocyanides that do not fall into either the Xanthocillin or Cyclopentyl classes are discussed below.
Mannitol-containing isocyanides, A32390A and brassicicolin A: 1,6-di-O-92- isocyano-3-methylcrotonyl-mannitol or A32390A is a novel antibiotic and antifungal agent isolated from the culture of the fungus Pyrenochaeta sp. A32390A is effective against the pathogenic yeast Candida albicans, the causative agent of the Candidiadis infection in humans 159,160. A32390A is also a dopamine beta hydroxylase (DBH)
161 inhibitor with an IC50 value of 1.7µg/ml . Like several other molecular targets of isocyanide natural products, DBH is a copper-containing oxygenase. DBH catalyzes the conversion of dopamine and other phenylethylamine derivatives to the neurotransmitter 31 norepinephrine using ascorbic acid as a cofactor 162,163. Deficiency of DBH results in impaired synthesis of catecholamines such as norepinephrine and epinephrine in the central and peripheral nervous system with a concurrent increase in serum dopamine levels 164. This condition of increased dopamine/norepinephrine ratio is known as
“norepinephrine deficiency” and it affects the autonomic nervous system. Physiological effects of norepinephrine deficiency include perturbed blood pressure regulation, elevated serum dopamine levels, and certain mental disorders 165–168. When injected intraperitoneally or subcutaneously, A32390A reduces heart and adrenal norepinephrine levels with a concurrent decline in blood pressure without any hepatotoxicity in DOCA hypertensive rats. It does not affect brain norepinephrine levels when injected and is ineffective when administered orally 161,169–171. Oral administration of isocyanides is likely to be generally problematic, as the isocyanide moiety is acid-labile and unlikely to survive passage through the stomach. Like most other isocyanides, the mechanism of
A32390A inhibition of DBH is unknown but likely involves interaction of the isocyanide moiety with the active site copper ion of DBH. Research is also required regarding the potential effect of A32390A in the induction of the mentioned disorders.
Another structurally similar isocyanide with a d-mannitol core is brassicicolin A, isolated from the plant necrotrophic fungus Alternaria brassicicola172. It is a potent antibiotic against Bacillus subtilis and Staphylococcus aureus 173. Interestingly, brassicicolin A is a phytotoxin (up to a concentration of 5mM) and a phytoalexin, which is a defensive chemical produced by plants as a defense mechanism against pathogenic infections, produced by infected Brassica juncea plants, unlike the other isocyanides discussed so far 174,175. 32
Indole-containing isocyanides: B371 is a novel indole containing isocyanide antibiotic isolated from Pseudomonas sp. 176. A similar isocyano indole metabolite, brasilidine A, was isolated from Actinomycete Nocardia brasiliensis 123 and possesses significant cytotoxicity against various tumor and multidrug resistance cell lines124.
Hazimicin factors 5,6: The hazimicins are a novel class of isocyanide antibiotics
(hazimicin factors 5 and 6), isolated from Micromonospora echinospora var challisensis
SCC 1411 177 ; they are isomers that are interconvertible in presence of water 2,21,178.
Aerocyanidine and Aerocavin: Aerocyanidine, an isocyanide antibiotic primarily effective against gram positive bacteria, was isolated from cultures of Chromobacterium violaceum ATCC 53434 179. In the same year, another isocyanide anttibiotic called aerocavin was isolated from the same bacterium and is effective against both gram positive and negative bacteria180.
1.1.2 Non-terrestrial isocyanides: All isocyanide natural products produced by marine organisms are terpenoids (isoprenoids), and maybe diterpenoids (4 isoprene units) or sesquiterpenoids (3 isoprene units). The first marine isocyanide to be discovered was axisonitrile 1, a sesquiterpenoid, from the sponge Axinella cannabina 20. This portion of the review will focus primarily on the types of marine isocyanides and their biological roles, biosynthesis and biochemical targets.
1.1.2.A Types of marine isocyanides: Depending on the number of isoprene units in their structures, marine based isocyanides can be classified into monoterpenoids, diterpenoids and sesquiterpenoids. This next section will briefly cover the different types of marine isocyanides and their biological roles. A more in depth and comprehensive coverage of this topic is available in previously published reviews 2,19,124,181,182. 33 i. Monoterpenoid marine isocyanides: This group mainly consists of monoterpenoid indole containing isocyanide alkaloids, synthesized primarily by blue green algae of the stigonemataceae family. Hapalindole A was the first such monoterpenoid indole isocyanide to be discovered from the cyanophyte Hapalosiphon fontinalis (Ag.) Bornet
192. This was followed by the discovery of eighteen additional isocyanide functionalized indole alkaloids from the same cyanophyte, named hapalindoles C-Q and T-V 183. The monoterpenoid type isocyano alkaloids can be classified into three primary groups124:
(a) the C21 indole alkaloids, which includes the hapalindoles, maybe further classified into tetracyclic and tricyclic depending on the presence or absence of a C4-C16 bond.
Other C21 type of isocyanides are the fischerindoles, which are basically tetracyclic hapalindoles modified to possess a C2-C16 bond rather than the C4-C16 bond.
(b) the oxindoles and the oxygenated C21 indoles which include the hapaloxindoles, the hapalonamides, the hapalindolinones, and the welwitindoles.
(c) the C26 indole alkaloids which include the ambiguines.
A thorough account of these indole isocyano alkaloids can be found in the reviews published by C.W.J Chang124 and by Bhat et al 185. These monoterpenoid isocyanide natural products are bioactive agents, primarily exhibiting antimicrobial properties, just like the terrestrial isocyanides40,185–187. Most of these natural metabolites are antifungal agents41,188,189; Ambiguines A-F and hapalindoles G and H were isolated from Fischerella ambigua 190 185; Ambiguines H and I exhibit both antifungal as well as antibacterial properties with MIC values comparable to streptomycin, puromycin and amphotericin41. Ambiguine isocyanides A, K and M show potent inhibitory activity against M. tuberculosis and B. anthracis 40. The terrestrial cyanobacterium Fischerella 34 sp is responsible for the synthesis of several isocyanide alkaloids which exhibit potent antimicrobial activities, such as Hapalindole A and Fischerindole L isolated from the cyanophyte Ficherella muscicola, shown to inhibit the growth of A. oryzae, P. notatum,
S. cerevisiae etc 191. Hapalindole T, isolated from the cyanobacterium Fischerella sp that grows in the bark of the Neem tree, is active against multiple pathogenic bacterial species such as M. tuberculosis, S. aureus, P aeruginosa, S. typhi, E. aerogens, E. coli, etc 192.
Hapalindoles A-H inhibit the growth primarily of pathogenic gram positive bacteria such as, Streptococcus sp (S. pneumoniae, S. faecium, S. pyogenes, S. epidermidis etc) 185.
Fischerella ambigua UTEX 1903 synthesizes several hapalindole type isocyanides, namely fischambiguines A and B, ambiguines P, Q and G, out of which fischambiguine
B shows strong inhibitory activity against M. tuberculosis 40. The welwitindolinone isocyanide alkaloids, including welwitindolinone A isocyanide, are known for their antifungal and anti-larval properties and are produced principally by the blue green algae
Hapalosiphon welwitschii 38. The welwitindolinones are not the only isocyanide natural products with insecticidal activities; Hapalindole L, 12- epi hapalindole C, 12-epi hapalindole E and 12-epi hapalindole J all kill Chironomus riparius larvae 185,193. A more detailed account of the antimicrobial activity of this class of isocyanides is discussed by
Bhat et al 185. Surprisingly, some of these alkaloids are also phytotoxic185. Matsumoto et al isolated ambiguine D from Hapalosiphon sp, which inhibited the growth of plants via an indirect inhibition of mitosis. This mitotic block was created by the accumulation of reactive oxygen species (ROS), mainly superoxide anion, generated via ROS-mediated lipid peroxidation 194–196. 35
The fact that most of these isocyanide alkaloids inhibit the growth of pathogenic microbes is a sharp contrast to some of the previously discussed isocyanides which potentially help in promoting pathogenicity and survival within the hosts.
Welwitindolinones and ambiguines are isocyanides that are synthesized by two separate gene operons, well and amb respectively (the well operon encodes well1-3 proteins which are homologs of the amb products amb1-3, mentioned previously). Both the well and the amb operon proteins are homologs of the pvcA-D proteins in P. aeruginosa. Most of the pvcA-D products (including the two siderohphores pyoverdine and pyochelin, the isocyanide paerucumarin, and exotoxins) have been implicated in the promotion of pathogenicity and survival within hosts, whereas the homologous welwitindolinones and the ambiguines products kill pathogenic bacteria and fungi rather than promote their survival.
Interestingly, welwitindolinones and their analogues have been shown to reverse
P glycoprotein-mediated multiple drug resistance (MDR)124,197–199 in human ovarian adenocarcinoma cell line (SK-OV 3), thereby preventing the expulsion of anti-cancer drugs. More details about the ability of welwitindolinones and their derivatives to reverse
MDR can be found in the comprehensive review by Bhat et al 185. Most of the hapalindole-type antibiotics possess anti-cancer properties as well. Kim et al 187 reported the cytotoxicity of several hapalindoles against different human cancer cell lines, including HT 29 (colon adenocarcinoma), MCF 7(breast carcinoma), NCI-H460 (large cell lung carcinoma), SF268 (glioblastoma) and IMR90 (lung fibroblast normal cell) cell lines. The mechanism of inhibition of cancer cell proliferation by hapalindoles remains unclear, although it might be a combination of different mechanisms including reversal of 36
MDR in drug-sensitive cancers. Another potential mechanism might be via inhibition of
RNA polymerase. Evidence to support this conjecture is provided 12 epi hapalindole E from Fischerella sp, which acts as an inhibitor of mRNA biosynthesis 200 by targeting
RNA polymerase. The consequences of inhibition of RNA polymerase are manifold, including the treatment of African trypanosomiasis 201 and the activation of p53- mediated apoptosis of cancer cells 202–205. Hapalindoles are also antagonists of the antidiuretic hormone arginine vasopressin; hapalindole A inhibits vassopressin binding to the kidney as well as inhibiting kidney vasopressin-stimulated adenylate cyclase185,206,207.
12 epi hapalindoles can be biosynthesized via two pathways185 : L-tryptophan reacts with geranyl pyrophosphate to form a terpene ((3-Z-20-isocyanoethyl indole) which can either undergo cyclization with geranyl pyrophosphate or undergo a chloronium ion mediated reaction with Z-3,7-dimethyl-1,3,6 octatriene to synthesize the chlorinated hapalindoles (such as 12-epi hapalindole E isocyanide). 12 epi hapalindole E is the precursor for all chlorinated isocyanide metabolites from H. welwitschii; details of biogenesis of these indole alkaloids can be found in Moore and Stratmann et al’s study 38 as well as in the review by Richter et al 208. The indole groups of hapalindole alkaloids are installed by the indolyl vinyl isocyanide synthases (Ambl1-3 and WelI1-3) using the substrates L- tryptophan and ribulose-5-phosphate 209. Welwitindolinones and ambiguines have operons (well and amb respectively) which encode isocyanide synthase
(isnA) and oxygenase (isnB) homologs (this has been covered in detail previously). A thorough description of the biological and chemical synthesis of these monoterpenoid alkaloids can be found in published reviews 185 124. 37 ii. Diterpenoid marine isocyanides: The first reported diterpenoid type isocyanide was
3- isocyano-3,7,11,15-tetramethylhexadeca-1,6,10,14-tertaene 210 from the marine sponge
Halichondria sp, almost a decade before the discovery of the previously described monoterpenoids. There are three classes of isocyanide diterpenoids:
(a) The acyclic diterpenoids: The first and the only observed acyclic isocyanide diterpenoid is 3- isocyano-3,7,11,15-tetramethylhexadeca-1,6,10,14-tertaene, isolated from a mixture of its corresponding isothiocyanate and formamide in Halichondria sp.
Out of all of these compounds which possess antimicrobial properties, this is the only isocyanide diterpenoid that is acyclic; all other diterpenoid iscocyanides are cyclic124,210.
(b) The kalihinanes: The second class of diterpenoid isocyanide natural products are the kalihinanes. These are bicyclic organic compounds which possess different types of tetrahydropyran (THP), tetrahydrofuran (THF) and dihydropyran (DHP) appendages attached to the carbon 7 position 124. Eleven different types of kalihinanes alcohols
(kalihinols) 124 were isolated from the sponge Acanthella sp by Patra and Chang et al
211,212. Kanihinols can also possess chloro, or alkene functional groups, and they constitute about 50% of all diterpenes isocyanides212. A thorough description of kalihinanes can be found in reviews published by Chang 124 and Garson and Simpson et al 181. All of the kalihinane isocyanides possess antimicrobial properties 213. Kalihinol A and F inhibit the growth of bacteria and fungi, such as, S. aureus, C. albicans, B. subtilis
2. Kalihinol A and two new isocyanide-containing diterpenoids, isokalihinol B and kalihinene, were isolated from the marine sponge Acanthella klethra. All three compounds are toxic to fungi such as Mortierella ramannianus and Penicillium chrysogenum. They are also cytotoxic to P388 murine leukemia cells 214. In addition, 38 some of these diterpenoids possess the ability to kill parasitic worms 215,216 and malarial parasites 217,218. Kalihinenes X,Y,Z, Kalihinol A and 10- formamidokalihinene from
Acanthella cavernosa exhibit antifouling activity against the larval settlement and metamorphosis of Balanud amphitrite 181,219,220.
(c) The amphilectanes: These are isocyanide-containing alcohols isolated from the marine sponge, Hymeniacidon amphilecta 221. All of the these diterpenoids are potent antimicrobial agents and may be further classified into the following types124: isocycloamphilectanes (primarily isolated from the marine sponge Cymbastela hooperi
222), cycloamphilectanes (primarily isolated from the sponges Amphimedon sp223,
Halichondria sp 224 , Adocidae sp 225 and Cymbastela hooperi 222) and neo- and pre- amphilectanes124. The last type of amphilectanes includes only two compounds, 7- isocyano isocycloamphilecta-11,15-diene from the sponge Adocidae sp 225 and 7- isocyano neoamphilecta-1,15-diene from Cymbastela hooperi 222. The isocycloamphilectane isocyanide, isocycloamphilectane-7,20-diisocyanoadociane from the sponge Amphimedon sp, 226 shows antimalarial activity181,222, along with several other marine isocyanide secondary metabolites 227,228. Although the mechanism of malaria prevention by these isocyanides is unclear, it may include the ability of isocyanides to bind iron porphyrins. This is consistent with the aforementioned ability of isocyanide natural products to act as siderophores. Additionally, isocycloamphilectane isocyanides may prevent conversion of hemozoin to beta hematin229,230; accumulation of the toxic heme then leads to disruption of the membranes of the malarial parasite181,231. iii. Sesquiterpenoid marine isocyanides: Sesquiterpenoids are principally isolated from marine sponges and molluscs, like the diterpenoids covered in the previous section. 39
Unlike the diterpenoids, sesquiterpenoids have only one nitrogenous moeity (-NC, -NCS,
-NHCHO, -SCN or -NCO). There are 105 of these sesquiterpenoids, of which only 35 are isocyanides while 35 others are isothiocyanates124. Sesquiterpenoids often are present with diterpenoids in the same organisms and have eight different skeleton types, discussed below124. One of the most prolific producers of sesquiterpenoids is the sponge
Axinella cannabina, which synthesizes about 33 different sesquiterpenoids, all of which belong to the P-U skeleton types. The different skeletal types are mentioned below:
(a) Axanes: These were the first marine isocyanide isoprenoid natural products discovered. The first two sesquiterpenoids bearing the axane skeleton to be isolated were axionitrile 1 and axisothiocyanate-1 from the sponge Axinella cannabina 20. Since then, several other axane natural products have been isolated from Axinella cannabina as well as from a different sponge, Acenthella cavernosa.
(b) Aromadendranes: The first aromadendrane type (Q type) sesquiterpenoids to be reported were axisonitrile 2 along with its isothiocyanate and amide derivatives, axisothiocyanate 2 and axamide 2 respectively from the sponge Axinella cannabina 232.
(c) Eudesmane: Acanthellin 1 was the first isocyanide sesquiterpenoid of the eudesmane type to be isolated from the sponge Acanthella acuta 233. Its corresponding isothiocyanate and formamides were isolated from a different sponge, A. cannabina 234.
(d) Epimaaliane: First isolated from the nudibranch Cadlina luteomarginata feeding on the sponge Axinella sp 235, these sesquiterpenoids are also found in the sponge A. cannabin.A detailed account of these compounds can be found in Chang’s review 124. 40
(e) Cadinane: Candinanes are synthesized from amorphene, (-) -10 isocyani-4- amorphene, and have been isolated from nudibranches (Phyllidia sp) as well as sponges
(Axinyssa sp, A. cannabina, A. pulcherrima)124.
(f) Spiroaxanes: These types of sesquiterpenoids were first isolated from the sponge
Axinella cannabina, the same sponge that synthesizes the axanes axionitrile 1 and axisothiocyanate-1. These three novel sesquiterpenoids possess an isocyano, insothiocyano, or amide groups and were named axisonitrile 3, axisothiocyanate 3 and axamide 3 respectively 236. Spiroaxane sesquiterpenoids have subsequently been isolated from several marine sponges, including Acanthella acuta237, A. klethra238, Axinyssa aplysinoides239, A. cavernosa 240,241, and somes nudibranches, such as Phyllidiia pustulosa and P. ocellate 240,242.
(g) Bisbolanes: The first sesquiterpenoid of this skeleton type was isolated from the sponge Theonella swinhoei 243. Of all known bisbolanes, only two have been identified to possess an isocyanide group 124, although isocyanates and formamides of this type are frequently found in nature.
(h) Pupukeananes: The first pupukeanane isocyanides were discovered from the nudibranch P. viscosa feeding on the sponge Hymeniacidon sp. The isolated isocyanides were 9-isocyanopupukeanane and its isomer 2-isocyanopupukeanane, both toxic to fishes
244,245.
Like most other isocyanide natural products, almost all sesquiterpenoid isocyanides are antimicrobial agents221,233,246–248. Metabolites of the marine sponge, Acanthella pulcherrima, including the isocyanide (-)-8 isocyano-7-aromadendrane, inhibit the growth of Bacillus subtilis 249. The sponge Axinella sp produces several isocyanide 41 sesquiterpenoids, including axisonitrile 2, which exhibit antimicrobial properties and protect the sponge from being eaten or overgrown248,250. Some of the sesquiterpenoids also exhibit antimalarial activities similar to the diterpenoids discussed previously.
Axisonitrile 3, its corresponding isothiocyanate, and three other eudesmane sesquiterpenoids show promise against Plasmodium falciparum, the causative agent of malaria 238.
Axisonitrile 2 and other sesquiterpenoid metabolites of the sponge Axinella spare antifouling agents; they prevent the settling and metamorphosis of the larvae of P. pacifica, S. tribranchiata, H. refescens in marine environments 248,250. Terpenoids from the sponge Acanthella cavernosa also show antifouling activity against the larvae of the barnacle Balanus amphitrite, as covered in the section detailing the kalihinanes241.
Nudibranches of the family Phyllidiidae synthesize isocyanide sesquiterpenoids, which also exhibit antifouling properties against larval attachment and metamorphosis of
Balanus Amphitrite242. Sesquiterpenoids are toxic to fishes as well; for example, axisonitrile 2 and other metabolites of Axinella sp are poisonous to goldfishes250, (-) 8- isocyano-7 aromadendrane and other metabolites from Acanthella sp are toxic to guppies
251,252. This toxicity may result from isocyanide-induced suppression of feeding in fishes250 by a mechanism that remains unclear. This isocyanide-induced toxicity of fishes may bolster the survival of marine sponges by killing off competition, as we have seen in the case of terrestrial isocyanides conferring survival advantage to isocyanide-producing microorganisms in the soil microbiome. Nudibranches typically feed on sponges that synthesize these piscicidal agents, allowing them to use these toxins as defense mechanisms235,245. Accounts detailing the chemical and biological synthesis of 42 isocyanide functionalized di- and sesqui- terpenoids already exist 151 2,124,181,182 and therefore won’t be covered in details in this review.
1.1.3 Degradation of isocyanides by Isocyanide hydratase (ICH)
Isocyanide hydratase (ICH) (EC 4.2.1.103) was isolated by the Kobayashi group from Pseudomonas putida N19-2 32. This was the first enzyme discovered that could convert a wide range of isocyanides into their corresponding formamides via addition of water to the isocyanide moiety. ICH is induced by the presence of isocyanides in the growth medium, confirming that isocyanide detoxification activity is not adventitious but is a specific response to isocyanides. Therefore, ICH plays a key role in the microbial defense against isocyanide natural products.
Further investigations into this enzyme revealed that it is a ~60 kDa homodimeric protein encoded by the 684-nucleotide long inhA gene. ICH is a cysteine-dependent enzyme whose active site Cys101 residue is essential for catalysis. Cys101 initiates nucleophilic attack at the isocyanide carbon resulting in the formation of a proposed thioimidate intermediate which is ultimately hydrolyzed to the corresponding formamide31,253. The thioimidate intermediate in ICH has not been previously observed using any technique, but it was assumed to be a mandatory intermediate formed by the nucleophilic attack of Cys 101 on the isocyanide carbon. The work described in this thesis will describe the first successful visualization of the thioimidate reaction intermediate during enzyme catalysis in microcrystals using serial femtosecond mix and inject crystallography.
Kobayashi et al isolated another protein with isocyanide hydratase activity from
Arthrobacter pascens F164. This second ICH enzyme is accompanied by a deformylase 43 that acts on isocyanide-derived formamides to give formate and amines 33. Therefore, this two-enzyme system suggests that Arthrobacter pascens F164 does not simply detoxif isocyanide natural products but can convert them to common metabolites with well- defined catabolic pathways. A comparison between the Pseudomonas putida N 19-2 ICH and the Arthrobacter pascens F164 ICH revealed no significant sequence similarity between the two as well as different structural, biochemical and immunological properties254.
Previously the Wilson group determined the crystal structure of the 48 kDa wildtype ICH isolated from Pseudomonas fluorescens at resolution 1.0 Å (PDB:
3NON)31. Site-directed mutagenesis in that study confirmed that Cys101 is catalytically essential, as mutations at Cys101 (C101S, C101A) resulted in inactive ICH. The active site Cys101 interacts with Asp17 and Thr102 in the ICH active site. The T102V mutation results in substrate inhibition, whereas the D17V, D17E and D17E mutations severely reduce the catalytic ability of ICH. Based on analysis of COOH bond lengths in these atomic resolution structures, Asp17 was shown to be protonated (neutral) and was proposed to activate an ordered water that hydrates the thioimidate intermediate. Another catalytically important residue is Arg214, which was reported to be important for anion binding near the active site by interacting with anions via its cationic side chain; suggesting that substrates for ICH may possess an anionic moiety. Interestingly, the ICH structure is similar to that of human DJ-1, however differences in the to the active sites of
ICH of DJ-1 render DJ-1 incapable of degrading isocyanides. Moreover, these variations in the ICH active site also disfavor oxidation of Cys101, whereas the active site of DJ-1 44 strongly favors oxidation of Cys106 to Cys106 -sulfinic acid, enabling DJ-1 to sense ROS levels cells.
The work described in this thesis clarified several outstanding mechanistic questions regarding ICH catalysis by directly observing catalytic turnover using serial mix-and-inject serial crystallography. In addition, this technique observed non- equilibrium motions in ICH that were activated by covalent modification of Cys101 that are functionally important for turnover.
45
Chapter 2: Introduction
The relationship between internal enzyme conformational dynamics and catalysis.
The role of protein conformational dynamics in enzyme catalysis has been a topic of much heated debate for several decades 255–258. The central question of this debate is: do enzyme dynamics contribute directly to transition state barrier crossing (the chemical event) or are they involved only in substrate binding and product release? Because the physiochemical basis of enzyme catalysis is still incompletely understood, internal protein motions and their contributions to effective substrate turnover has garnered much attention in the past two decades. Motions of amino acid residues, in or around the active sites, can have significant impact on substrate turnover. As such, the identification and characterization of such motions and their overall catalytic consequences has become of paramount significance259,260. This section of the thesis will describe some examples where internal dynamics in enzymes effect their overall catalysis258,261,262 and delineate mechanistically how such events affect passage along the reaction coordinate263.This topic is important for this thesis, which addresses how covalent modifications of the active site reactive cysteine 101 in Isocyanide Hydratase (ICH) enzyme triggers a charge- gated conformational switching of a nearby helix H. This catalysis-activated motion of helix H is important for efficient ICH catalysis. We propose that several enzymes may rely on internal active site dynamics for catalysis, particularly when triggered by active site modification. ICH is the first system, to our knowledge, where cysteine non- disulfide covalent modification directly gates conformational dynamics essential for its function.
46
2.1. How internal protein motions effect enzyme catalysis.
During a catalytic cycle, enzymes can undergo conformational changes during any of the key catalytic steps, including substrate binding, intermediate formation/resolution, product formation/release, etc. These motions, which occur on timescales ranging from milliseconds to seconds264, often define the rate limiting steps and thus the overall rates of the reactions258,265–269. A common way to estimate the degree of transition state stabilization by an enzyme is to compare the overall rates of the enzyme-catalyzed vs uncatalyzed reactions261. However, a limitation of this approach is that it provides a lower limit for transition state stabilization, as the rate-limiting step may not be the chemical step. One example of the interplay of enzyme conformational changes and catalytic rate is the enzyme Ribonuclease A (RNase A)270–272, which catalyzes cleavage of P–O5′ bonds in RNA272. The enzyme contains two mobile loops,
Loop1 and Loop4. These loops undergo a 2-3 Å conformational change upon substrate/ligand binding, resulting in switching from the “open” form (optimal for substrate / ligand binding) to the “closed” form to facilitate catalysis. This conformational switching, coupled to product release, is the rate limiting step of the reaction273–275,275–284.
However, whether this intrinsic conformational flexibility in RNase A aids catalysis by stabilizing the transition state remains unclear.
Transition state stabilization could potentially be affected by protein motions, providing a direct connection between enzyme dynamics and the mechanism(s) by which enzymes achieve their catalytic effects263,285–287. Transitions states are “balance points of catalysis” 288, when chemical bonds are being made/broken and the reaction has an equal probability of going in either direction; it is that fleeting unstable state where a molecule 47 is no longer a substrate but is not yet a product 289. All enzymes must stabilize the transition state for efficient catalysis16–23,23–27, or in other words, minimize the activation energy of the reaction (energy of the highly unstable transition state) to begin substrate turnover. Linus Pauling proposed that this likely occurs via selective, tight binding of the enzyme to the transition state species285. Specific mechanisms that enzymes use to stabilize the transition state include pre-organizing active site electrostatic environments302,303, and specific arrays of active site hydrogen bonds that favor binding to the transition state over the substrate 261,304. As an example that combines these strategies, Schowen et al’s minireview 305 describes transition state stabilization by proton bridging in enzymes involved in general acid-base catalysis306. Therefore, enzyme dynamics may be able to impart dynamic electronic microenvironments to active sites that are conducive to the formation/stabilization of the transition state species.307
The enzyme Dihydrofolate reductase (DHFR) has been proposed as an example of enzyme dynamics directly assisting transition barrier passage. E. coli DHFR enzyme possesses a highly conserved mobile loop308,309310, called the Met20 loop, whose mobility regulates access of substrate, solvent, and co-factor accessibility to active site pocket. The open conformation of the Met20 loop facilitates the binding of both NADP+ and folate. Upon binding, the disordered portion of the Met20 loop (residues 16- 20) undergoes a conformational change to form a hairpin turn that ensures proper alignment of the nicotinamide and pteridine rings in the active site310. This conformation also positions residues Ile14 and Met 20 near the active site they exclude water and stabilize the Michaelis complex via a network of hydrogen bonds and hydrophobic interactions.
To test the influence of Met20 loop mobility on E. coli DHFR catalysis, a mutant that 48 deleted four critical hairpin forming residues (16-19) from the Met20 loop (called DL1) was made. In wild type DHFR, product release is the rate limiting step311. In the DL1 E. coli DHFR mutant, hydride transfer becomes rate limiting (rate decreases from 950 sec-1 in wildtype to 1.7 sec-1 in DL1 mutant DHFR), arguing that the Met20loop is important for rate acceleration261,310.
It is possible that the mobile Met20 loop residues in E. coli DHFR play an important role in “compacting” the active site to correctly position the substrate and the co-factor and to reduce the total distance between donor and acceptor atoms to promote hydride transfer 312. Internal active site dynamics in DHFR supports a model of catalysis where the “open” active site can easily access and bind the substrate and co-factor, following which the disordered central portion of the Met20 loop adopts a hairpin turn which stabilizes the transition state310; inability to switch between open and hairpin conformations in mutant DL1 has a detrimental effect on catalysis310. As such, E. coli
DHFR has been subject to a number of combined experimental and computational studies to investigate the controversial role of enzyme dynamics in catalysis185,264,278,312–315.
Several of these studies by the Benkovic, Hammes-Schiffer, and Wright groups have proposed that collective motions in E. coli DHFR can serve as “promoting motions” that directly facilitate transition state barrier passage, although this is currently debated
288,316,317.
Other researchers have argued that, although such conformational dynamics can affect the overall rate of reaction or the rate limiting step of the reaction, they cannot contribute to catalysis via transition state stabilization or enhancing rates of barrier passage beyond what is permitted by equilibrium and the Eyring rate theory258,307,318–323. 49
One of the primary opponents of dynamical contributions to enzyme catalysis is Arieh
Warshel, who argues that in DHFR and other enzymes, catalysis is achieved by adoption of a specific electrostatic microenvironment in the enzyme’s active site, not by internal millisecond conformational dynamics317,324,325. Since most enzymatic reactions involve polar transition states, enzymes can decrease the activation energy barrier of the reaction by fashioning active site electrostatics to stabilize these transition states291,299,324,326,327.
Investigations into the connection between protein dynamics and the probability of reaching the transition state reveal a dominant role for the electrostatic fluctuations of the protein; if these electrostatic fluctuations differ in enzyme and in solution, they can contribute to catalysis321,328, otherwise not.
Reconciling the computational finding that dynamical events likely cannot contribute to catalysis with the results of studies on DHFR (and other enzymes) indicating that dynamics are important for catalysis is challenging. A plausible model is that dynamics are important for progress along the reaction coordinate, although they do not aid barrier crossing per se. In this thesis, we find that the enzyme Isocyanide
Hydratase (ICH) facilitates catalysis via changes in conformational ensembles that are gated by the formation of a catalytic intermediate. In ICH, the formation of a thioimidate catalytic intermediate eliminates negative charge on the active site cysteine thiolate that alters the dynamics of ICH, facilitating subsequent steps of catalysis.
A central concern when considering the connection between conformational dynamics and catalysis is correlating the timescales of these conformational dynamic events to key steps in catalysis. If such correlations exist, then the internal protein motions whose timescales correspond to key steps in catalysis are candidates for 50 functionally important motions. Generally, there are three types of protein motions. The first type is the local motion(s) (10−15 to 10−1sec) which include atomic fluctuations and side chain mobility. The second type of motions are rigid body motions (10−9 to 1 sec) which involve the movement of small parts of the protein; they include helix mobility, domain and subunit motions etc. The third and final type of motion is the large-scale slow motions (10−7 to 10 sec), which involve helix coil transformations, folding/unfolding etc.
Since the overall enzyme catalyzed reaction rate is related to the activation free energy barrier height, hypothetically one of the ways in which internal motions could potentially affect the catalytic rate is by decreasing the height of the activation energy barrier with the previously mentioned local (10−15 to 10−1sec) or rigid body motions (10−9 to 1 sec). In most cases, however, the enzyme transition state formation/existence is a femtosecond
(fs) timescale event. Therefore, these rigid body motions (conformational dynamics) are too slow to stabilize the transition state. In several enzymes, such as DHFR, Triose phosphate isomerase (TIM), Lactate dehydrogenase (LDH) and beta lactamase, the internal motions implicated in contributing to catalysis are of the rigid body type and therefore are too “slow” and cannot stabilize the transition state, but can potentially facilitate the later steps of catalysis.
Motions that involve substrate/ligand binding and organization of the active site electrostatic microenvironment to make it conducive for transition state binding, as well as internal dynamics that enhance populations of transition state accessible conformations, are slow, often in the scale of microseconds to milliseconds307,329. Faster fs motions occur after these slower protein motions and can stabilize the transition state to facilitate product release and to regenerate the native enzyme. However, there is no 51 evidence currently available to indicate that these fast motions are actually “directly linked to the bond breaking and forming process of the transition state”262.
Transition states have lifetimes on the timescale of single bond vibrational frequencies and as such, coupling of femtosecond (fs) bond vibrations to conformational motions of the millisecond (ms) scale is highly unlikely329. Warshel et al 330 have demonstrated that the kinetic energy of the conformational motion associated with substrate binding is entirely dissipated before the chemical step begins and therefore cannot affect it. Combined with computer simulations, they establish that in case of enzyme catalyzed reactions where the chemical barrier is greater than a few Kcal/mol, there exists no coupling between the chemical step(s) of catalysis and internal protein conformational dynamics.
52
Chapter 3: Introduction
Biochemistry of reactive cysteine residues in enzymes
3.1: The relationship between cysteine covalent modification gated enzyme dynamics and catalysis.
Although protein dynamics are not likely to contribute to barrier crossing directly, they can facilitate catalysis by promoting progress along the reaction coordinate. In this thesis, we explore how conformational dynamics can aid enzyme turnover using
Isocyanide Hydratase (ICH) as a model system. The present study involves reactive cysteine covalent modification(s) which alter the active site electrostatic microenvironment, resulting in helical dynamics important for efficient catalysis by ICH.
Having discussed the relationship between internal dynamics and enzyme catalysis in
Chapter 2, we will make an in-depth investigation into how protein dynamics can be coupled to cysteine covalent modification(s) in cysteine dependent enzymes in this section.
A multitude of post-translational protein modifications exist and are essential in mediating proper function for many proteins 331–334. Out of the 20 frequently occurring amino acid residues in human proteins, cysteine (Cys) is the most reactive due to its strongly nucleophilic thiolate side chain when ionized332. Cys residues are usually reactive, redox sensitive, polarizable and extremely sensitive to local microenvironment electrostatics335. Although cysteine makes up only about 2% 336 of protein amino acids in mammals, covalent modifications of cysteines have important consequences for protein function that affect essential processes such as cell growth, repair and death. As a result, 53 reactive cysteine residues are frequently located at functionally important sites in proteins and act as regulatory switches that can controls multiple downstream processes335,337.
Cysteines can be classified into several functional categories, such as catalytic, redox active, regulatory, structural, metal-coordinating, and some without any known role; a mini review detailing the description of these classes of cysteine residues is available by
Fomenko et al 337.
Cysteines that control downstream metabolic signaling processes in response to covalent modifications are widely studied337,338. These modifications arise naturally, as a consequence of reaction of cysteines with pools of reactive electrophilic metabolites in the cell 331,339,340. Some common cysteine covalent modifications include alkylation, oxidation, formation of metal (electrostatic) complexes, etc. Below I delineate the most common cysteine modifications and review examples of such modification(s) and their functional effects in cysteine dependent enzyme catalysis.
3.2.a) Unmodified cysteine (neutral thiol/negative thiolate).
An important regulator of the activity of cysteine-dependent enzymes is the redox state of their immediate microenvironment, which can be altered by the generation of reactive oxygen species (ROS)341. The d-orbitals of the sulfur atom of the thiol group allows for numerous oxidation states, permitting a multitude of possible oxidative posttranslational modifications. Reactive oxygen and nitrogen species can participate in signal transduction pathways via the reversible or irreversible oxidation of cysteine thiol groups in redox sensitive proteins, including several cysteine-dependent enzymes
342,343,92–94. The anionic sulfur atom in cysteine thiolates (R-S-) is the reactive form and is susceptible to oxidation by a large variety of ROS. Cys can be oxidized to various 54 species, discussed in detail later. Therefore, cysteines impart oxidant sensitivity to a multitude of proteins, with diverse regulatory and signaling consequences 347,348. There are several examples of cysteine-dependent enzymes of biomedical relevance that have redox sensing as a primary and secondary activity347. Reversible modifications are often held to be the most valuable for signaling, and both cysteine sulfenic acid and disulfides are reversible oxidative modifications that cause active site structural and functional changes in proteins 349, allowing thiols to act as redox regulatory switches350.
Figure 1: Major oxidation states of cysteines
The reactive form of thiols (R-SH) is the deprotonated form; the negatively charged
- 346 thiolate ion (R-S ) . Therefore, thiol pKa value is commonly used to determine if a particular cysteine residue is likely to be reactive. Because the pKa of an unperturbed cysteine residue is ~8.5, the electrostatic microenvironment of the active site can easily promote ionization of a cysteine to the thiolate in the physiological pH range. For cysteine-dependent enzymes, this pKa depression is important for nucleophilic catalysis.351. In the work described in this thesis, ICH undergoes active site conformational switching in response to catalytic cysteine modification, which changes the active site electrostatic microenvironment. As noted above, the ability to manipulate 55 the reaction buffer pH to control the extent of reactive cysteine protonation state provided us with a useful tool to fine tune the internal CH motions. This portion of the work is explained in more detail in Chapter 4 of this thesis.
3.2.b) Oxidative cysteine modifications.
As discussed in the previous section, cysteines can act as redox sensing switches via their ionizable thiol groups. As such, cysteine-dependent enzymes can play important roles in redox regulation and diseases with a redox component.
Oxidative phosphorylation is the process by which ATP is synthesized in the mitochondria by the reduction of molecular oxygen to water through a series of electron transfer events. However, during this process, a small portion of the oxygen does not convert to water, but is incompletely reduced to generate reactive oxygen and nitrogen species (ROS/RNS) such as peroxides, superoxides, hydroxyl radicals, peroxynitrite etc.
352, 353, 354. This induces oxidative stress by perturbing the normal redox state of the cells and can damage DNA, proteins and lipids, impair cell signaling, energy metabolism, and cause overall dysfunction of biological activity 355. During oxidative stress, increased levels of ROS/RNS target cysteine thiols for reversible or irreversible oxidation or glutathionylation338,356–359 and affect redox sensing and signaling. Fig. 1 summarizes the major oxidation forms of cysteines. As mentioned above, reversible oxidized species of thiols with known redox signaling roles are Cys-sulfenic acid and intra/intermolecular disulfide bonds, including glutathionylation349,360–362.
A) Disulfide formation: Disulfide bonds, or disulfide bridges or S-S bonds (R-S-S-R) are reversible covalent bonds that form between two proximal cysteine thiols within or 56 between individual polypeptide chains, by the oxidation of sulfhydryl groups under mildly oxidizing conditions359,363–365 (Reaction I).
2 R-SH→ R-S-S-R +2 H+ +2 e- …………………………. (I)
Disulfide bonds can also form from pre-existing disulfide bonds by the exchange of a thiol group (R-SH) with the disulfide (R’-S-S-R’) in a reaction called “thiol-disulfide exchange” or “disulfide scrambling”, catalyzed by various oxidoreductase enzymes 366–
368. This results in the generation of a new thiol group (R’-SH) containing cysteine and a new disulfide (R-S-S-R’)369–371 (Reaction II). This is often the rate determining step in the folding process of certain proteins344 and can occur in several cellular compartments incluing the ER, the mitochondrial inner membrane space (IMS) in eukaryotes, and the periplasm in prokaryotes344,372–374.
R′-S-S-R′ + R-SH → R-S-S-R′ + R′-SH …………………. (II)
The comprehensive review by Ziegler depicts a complete list of enzymes whose
375 functions rely upon disulfide-thiol exchanges . Elevated H2O2 levels in E. coli cause the cysteine-dependent OxyR transcription factor to upregulate the expression of antioxidant genes such as oxyS, katG and gorA to combat the effects of hydrogen peroxide.376. OxyR is activated by a conformational change resulting from the formation of an intramolecular disulfide bond between the cysteines Cys199 and Cys208376. OxyR is one of the best- studied examples of disulfide bond-mediated regulation of the cellular antioxidant response during oxidative stress. Disulfide bond formation can also promote bacterial pathogenicity and virulence377. While formation of S-S bonds confers structural stability to the enzyme, it can also act as a redox sensitive regulatory switch (breaking and remaking of disulfide bonds can regulate enzyme function)364,368,378–381. Moreover, the 57 ratio of cysteine to the disulfide (RS-SR)/cystine (CysSSCys) form has been shown to be dysregulated in several diseases in humans, including atherosclerosis379,382,383 and cancer384–387. Some examples of established disulfide bond regulation349 include activation of tissue factor (TF)388, redox sensing and allosteric monitoring in human carbamoyl phosphate synthetase389, redox sensitive global regulator in Staph. Aureus390, and allosteric control of S1A serine proteases391. Formation of disulfide bonds depends on three key factors392; the availability of a proximal partner cysteine with correct geometry for disulfide formation, the pH of the local microenvironment (pH> pKa of the cysteine thiols favors reactivity) and the redox state of the local microenvironment
(oxidizing conditions favor S-S bond formation). Since the likelihood of disulfide bond formation is strongly influenced by local cellular conditions, specific cellular compartments have evolved in both prokaryotes and eukaryotes to support this phenomenon. In bacteria, disulfide bond formation mostly occurs in the periplasm393, while in eukaryotes, disulfide bond breaking and making is most common in the endoplasmic reticulum (ER)392,394,395. The cytosol, being a more reducing environment compared to the ER, rarely facilitates disulfide bond formation, although there are exceptions. We discuss one such exception in Chapter 5, where the Isocyanide Hydratase
(ICH) enzyme from Ralstonia sp. contains two non-catalytic cysteines, Cys147 and
Cys220 which form disulfide bonds despite ICH being a non-ER protein.
Protein disulfide isomerase (PDI) is a family of endoplasmic reticulum (ER) enzymes that oxidize cysteine thiols, which then form disulfides with other nearby cysteines364,373,396–398. Extensive reviews have covered the topic of formation of disulfide bonds, the enzymes involved, and their mechanisms in both prokaryotes and 58 eukaryotes364,373,399–403. Even though the biochemistry of disulfide bond formation is well described, there is comparatively little known about the process of regulated disulfide bond reduction in the ER of mammals. Currently, there are no known PDI reductases or a glutathione reductases in the ER to reduce the disulfide bonds364,404. In contrast to mammals, the bacterial periplasmic reductive mechanism of disulfide breaking is very well characterized364,405–407.
B) Sulfenic, Sulfinic, Sulfonic acid formation: Oxidation of the sulfur atom in cysteine results in the formation of a series of organosulfur acids with progressively increasing oxidation states; they are cysteine sulfenic, sulfinic and sulfonic acids respectively408–410
(Fig. 1).
Sulfenic acids are reactive, unstable oxyacids generated as a result of oxidation of
- 359,410–414 cysteine thiols (R-SH/RS ) with oxidants such as H2O2 , alkylhydroperoxides and peroxynitrites359,411. They may also form as a result of oxidation of the cysteine with molecular oxygen, catalyzed by the cysteine dioxygenase enzyme415. These transient compounds can exhibit both electrophilic as well as nucleophilic characteristics416; they can react with neighboring thiols to form disulfides359,416,417 (Reaction III) or with nitrogen nucleophiles to generate the rarer sulfenamides410,418–421. Sulfenic acids (R-SOH) can be converted back to thiol (R-SH) by Thioredoxins (Trx)410 or other disulfide-based reduction strategies involving transient glutathionylation followed by reduction. In the E. coli periplasm, several proteins possess cysteine residues which form disulfide bonds. In periplasmic proteins with single cysteines (lacking disulfide bond forming partner), these residues are protected against oxidation to sulfenic, sulfinic and sulfonic acids by two thioredoxin related proteins, DsbG and DsbC422. 59
Sulfenic acids may also self-condense to form thiosulfinates359,423 (Reaction IV).
Since these sulfenic acid derivatives are also unstable, they have proven difficult to isolate and study359,424,425.
2 R-SOH + R-SH→ R-S-S-R +H2O ………………………….
(III)
2 R-SOH→ R-S(O)-S-R +H2O ……………………….
(IV)
Table 1 of Lo Conte and Carroll’s review 410 contains a list of biologically important proteins that form cysteine sulfenic acids and the authors discuss the regulatory implication of some of these sulfenic acids in cellular signaling 426–429. Protein tyrosine phosphatases420,421,430, cysteine proteases, papain, and glyceraldehyde-3- phosphate347,359,410,425, NADH oxidases and peroxidases, peroxiredoxins431, glutathione reductases, and human serum albumin (HA) are all regulated by sulfenylation412,432,433.
Other examples include the reversible oxidation of key cysteine-S- residues in transcription factors Fos and Jun (activator protein 1) and bovine papillomavirus1 E2 protein, which inhibits DNA binding in response to intracellular redox sensing359,425.
Sulfenic acid formation also helps in CD8+ T cell activation and proliferation434. Roos and Messens’ review 435 presents some of the most important biological roles of sulfenic acids in living systems.
Sulfenic acids can be further oxidized to more stable sulfinic acids (R-SO2H)
(Table 2 of Lo Conte and Carroll’s review 410). Some examples of biologically relevant proteins whose functions are mediated by formation of sulfinic acids, include peroxiredoxins359,436–438, peroxidase (Prx)437, DJ-1 439–442, and hydratases (Nitrile 60
Hydratase, SCNase) 410,443,444. Inactivation of peroxiredoxins by the formation of Cys- sulfinic acids438 can be reversed by the ATP-dependent enzyme Sulfiredoxin445–450, 451,452.
Sulfinic acids can be further oxidized to form the most oxidized form of cysteine in proteins, sulfonic acid (R-SO3H). An example of possible biological relevance of cysteine sulfonic acids is found in the peroxiredoxins, which converts to a dodecameric oligomer that exhibits enhanced chaperone activity when the active site cysteine to oxidized to the sulfonate, although the relevance of this activity is debated453,454.
Protein tyrosine phosphatases (PTPs) catalyze the dephosphorylation of phosphotyrosine in proteins455. PTPs are redox regulated by oxidation of the catalytic cysteine, which abrogrates enzyme activity456–458. While sulfenic acid can be easily reduced, reduction of the more stable sulfinic acid requires the dedicated sulfiredoxin enzyme and sulfonic acids cannot be reduced under intercellular conditions421. The oxidized cysteine in PTP, however, is capable of forming a sulfenyl-amide (-SN-) bond with the nitrogen atom in the polypeptide backbone, thereby protecting the cysteine from further oxidation455,459. Protein tyrosine kinase (PTK) can also be regulated by ROS and oxidative stress conditions. PTKs in the Src and Fibroblast Growth Factor Receptor
(FGFR) families are inactivated in response to oxidation of their reactive cysteine
(Cys277)460. The epidermal Growth Factor Receptor (EGFR) family of receptor tyrosine kinases mediate several important biological functions in the body, such as growth, differentiation, proliferation, protection from cancers. Activation of EGFR results in generation of ROS (H2O2), which in turn, oxidizes cysteine residues within redox sensitive proteins. Moreover, EGFR itself is regulated via oxidation of its active site
Cys797461. 61
The thioredoxin (TRX) and Nucleoredoxin (NRX) family of proteins undergo similar redox regulation as PTPs, by virtue of their reactive cysteines455. TRX is the archetypal thiol-disulfide oxidoreductase whose reversible disulfide formation is essential for its catalytic activity as a thiol reductant. In addition to its primary role in maintaining redox homeostasis, the oxidation of the cysteine pair 462,463 in TRX to form intramolecular disulfide bonds causes its disassociation from the enzyme Apoptosis signal regulating kinase 1 (ASK1)349. TRX is an inhibitor of ASK1, and its oxidation- regulated dissociation activates ASK1 to induce ROS and apoptosis455,459,464,465. NRX negatively regulates Wnt signaling by binding to and inhibiting DvI (dishevelled), which is an essential adaptor protein involved in Wnt signaling 466. When the catalytic cysteine pair in NRX is oxidized, binding to DvI is abolished and Wnt signaling is promoted,; therefore, NRX plays a redox responsive role in mediating DvI activity similar to the
TRX/ASK1 interaction455,467–470.
Another protein family with prominent roles in intracellular redox signaling is the peroxiredoxin (PRX) family of proteins. These proteins reduce hydrogen peroxide to water simultaneously with the oxidation of a reactive cysteine (peroxidatic cysteine, CP) to sulfenic acid (CP-SOH), which then reacts with another cysteine (resolving cysteine,
471 CR) to form a disulfide bond . Depending on the presence or absence of the CR, PXS enzymes can be classified into 2-Cys or 1-Cys PRX subfamilies472–474. Oxidized 2-Cys
PRX is then reduced and recycled by the TRX system and 1-Cys PRXs by glutaredoxin
474–476. Under conditions of high oxidative stress, PRX forms homo oligomers which bind and activate mammalian Ste20 like kinase 1 (MST1), leading to cell death and apoptosis455,477–479. Irreversible oxidation of PRX cysteines into the hyperoxidized 62 sulfinic and sulfonic acid forms inactivates PRX and serves as a cellular marker for oxidative damage454.
Cysteine oxidation has consequences in energy metabolism as well.
Glyceraldehyde-3-phosphate dehydrogenase (GAPDH), a critical glycolytic enzyme, can be oxidized at its active site cysteine residue to form a sulfenic acid or mixed disulfides with glutathione or other proteins. This oxidation prevents glycolysis and perturbs energy homeostasis and glucose metabolism349,480,481. Therefore, proteins containing reactive cysteine residues can be modified by a range of electrophilic species produced by metabolism or stress and thus can participate in multiple biochemical and signaling pathways.
C) Other thiol oxidation processes: In addition to ROS, cysteine thiols (R-SH) are oxidized by glutathione (S- glutathionylation: S-SG) and nitric oxide (S- nitrosylation: S-
NO) (Fig. 1).
S- nitrosylation is the process of covalent attachment of nitric oxide (NO) group to a cysteine thiol (R-SH) group to form a cysteine-nitosothiol (R-SNO) 482–484.
Nitrosylation has emerged as an important redox regulatory mechanism for mediating cGMP-independent NO signaling effects 485–487 as well as for combating nitrosative stress486,488.
Cys-nitrosothiols (R-SNO) are involved in multiple biological processes489, including apoptosis447, cardiovascular signaling485,490, combating cold stress in plants491,
HIV-1 protease inactivation492, transient receptor potential (TRP) channel activation493, ion channel regulation, signal transduction, neurotransmission, DNA repair, etc489. In protein tyrosine phosphatase 1B (PTP1B), S-nitrosylation inhibts irreversible oxidation 63 of the catalytic cysteine (Cys215) into sulfinic and sulfonic acid species, thereby protecting the enzyme from irreparable oxidative damage487. S-nitrosylation has also been implicated in defense against microbial infections and cancer494. An excellent account of the biological functions of cysteine nitrosothiols is available in Broillet’s extensive review489.
Glutathionylation of cysteines is another mechanism by which cysteine- containing proteins/enzymes act as redox signal transduction switches495,496. Cysteine sufenic and sulfinic acids can be enzymatically (glutathione-s-transferases, glutaredoxins) or non-enzymatically reduced and conjugated to glutathione (GSH) to form S- glutathionylated cysteines497 (Fig.1). Under conditions of nitrosative stress, cysteines can be nitrosylated, as discussed previously; the nitrosothiols can then be converted into S- glutathionylated cysteines with or without S-nitrosoglutathione (GSNO) as a reaction intermediate497 (reactions V-VII).
PS-N = O+ GSH→ P-S-S-G +HNO …………………………. (V)
PS-N = O+ GSH→ PSH + GSNO …………………………. (VI)
PSH + GSNO → P-S-S-G +HNO …………………………. (VII)
Inside a cell, the ratio of reduced glutathione (GSH) to oxidized glutathione (GSSG) is known as the GSH/GSSG ratio which informs on the redox capacity or balance of the cell. This balance is kept in check by redox reactions catalyzed by oxidoreductases such as GSH peroxidase and GSH reductase. Disturbance of the GSH/GSSG ratio by increased concentrations of ROS and RNS results in several pathological conditions such as apoptosis and necrosis497. Proteins that undergo S-glutathionylation are typically cytoskeletal, glycolytic, or involved in energy metabolism, signaling pathways, calcium 64 homeostasis etc497. Some of the signaling pathways impacted by glutathionylation of reactive cysteines include mitogenic signaling pathways (Ras MEK ERK pathway, PTPs,
Protein kinase A)498–501, PI3K signaling498,502–505, I κB kinase-nuclear factor κB pathway497,506–508, pro apoptotic JNK-c-Jun pathway in cancer cells497,506,509–517.
Townsend et al. provide a detailed review of the biological and physiological mechanistic impact of cysteine glutathionylation497. Redox regulation of mitochondrial functions by glutathionylation of mitochondrial proteins and their effect on energy metabolism is comprehensively reviewed by Mailloux et al518.
3.2.c) Other cysteine modifications:
Other cysteine modifications include metal-mediated modifications and desulfurization (removal of cysteine thiol, including oxidative elimination of cysteine to form dehydroalanine (DHA)519. These type of cysteine modifications are reviewed by
Chalker et al 65.
3.3. Cysteine covalent modification and internal enzyme dynamics:
Covalent modification is a common and physiologically important perturbation to proteins. Reactive residues such as cysteine are prone to diverse covalent modifications with catalytic or regulatory consequences. In the work described in this thesis, charge neutralization of the reactive Cys101thiolate in Isocyanide hydratase (ICH) enzyme triggers active site helical dynamics important for efficient substrate catalysis. The cysteine modification in ICH remodels active site H-bonding networks and gates catalytically important changes in protein dynamics with a clearly defined mechanism. It is very common for protein structures to fluctuate owing to thermal motion and in response to functional changes such as ligand binding. As a consequence, it is 65 challenging to determine which protein motions are functionally most important at equilibrium. Enzymes that are transiently and covalently modified during catalysis offer a way to identify functional motions, as the modification can trigger catalytically important conformational changes. The covalent modification of the active-site cysteine in isocyanide hydratase weakens a critical hydrogen bond required for reactivity. Hydrogen bond disruption triggers a cascade of conformational changes whose modulation by mutation is detrimental to enzyme turnover. Most enzymes that form catalytic intermediates will experience similar transient changes in active-site electrostatics, suggesting that modification-gated conformational dynamics is common in enzymes.
Notably, all cysteine thiolates will experience a similar loss of negative charge upon covalent bond formation. Because cysteine residues can have multiple roles in a protein, including acting as a catalytic nucleophile, metal ligand, acylation target, redox target, and others, many different modification-mediated signals may be transduced through altered cysteine electrostatics to impact protein dynamics and function. Thus, cysteine can potentially significantly expand the ways in which protein biophysical properties are coupled or respond to cellular needs.
Enzymes catalyze biochemical reactions by binding, conformationally organizing, and lowering the activation energy barrier of reactants 520. Aside from accelerating certain biochemical reactions, enzymes also inhibit undesired, off-pathway reactions involving highly reactive intermediate species521. The collective dynamics of enzyme and substrate are often tightly coupled to the catalytic cycle 521–525. For example, substrate binding can shift the enzyme’s conformational distribution to structurally and electrostatically re-organize the active site. More controversially, the coupling of 66 conformational dynamics and electrostatics has been proposed to directly contribute to the chemical step of enzyme catalysis, although their precise roles remain actively debated256,264,523,526–529 . Formation of catalytic intermediates transiently modifies the enzyme and may lead to a new protein motion regime that facilitates later steps of catalysis. These motions often span vastly different time-scales, from fs-ns for barrier crossing to ms or longer for product release. While functionally important motions may be present even in a resting enzyme530 , distinguishing functional motions from the many other fluctuations of a macromolecule at equilibrium is challenging523, leaving major questions about the role of dynamics in catalysis unanswered. For example, how do catalytic intermediates reorganize an enzyme’s conformational landscape to facilitate later steps in the catalytic cycle? Integrating recent developments in serial and conventional RT crystallography with advanced computational methods531 can offer new, exciting opportunities to reveal enzyme mechanism and interrogate the consequences of cysteine modification on protein dynamics.
67
CHAPTER 4:
Mix and Inject Serial Femtosecond Crystallography at XFEL reveals gated
conformational dynamics during ICH catalysis.
Note: The results described in this chapter have been published, all text is modified from the
original version “Mix-and-inject XFEL crystallography reveals gated conformational dynamics
during enzyme catalysis”.
https://doi.org/10.1073/pnas.1901864116
The catalytic cycle of ICH. Isocyanide substrate binds with ICH Cys101-S− poised for nucleophilic attack and with the Cys101-Ile152 H-bond intact. Formation of the thioimidate intermediate weakens the Cys101-Ile152 H-bond and causes helical motion (magenta helices H and I) that promotes intermediate hydrolysis (red sphere; curved arrow). Hydrolysis of the thioimidate intermediate restores the reactive C101-S− and strengthens the Cys101-Ile152 H-bond, favoring the strained helical conformation.
68
4.1 Abstract:
How changes in enzyme structure and dynamics facilitate passage along the reaction coordinate in catalysis is a fundamental, unanswered question. Here, we use time-resolved, Mix-and-Inject Serial Crystallography (MISC) at an X-ray Free Electron
Laser (XFEL), ambient temperature X-ray crystallography, mutagenesis and enzyme kinetics to characterize how covalent catalysis modulates isocyanide hydratase (ICH) conformational dynamics throughout its catalytic cycle. We visualize this previously hypothetical reaction mechanism, directly observing formation of a thioimidate covalent intermediate in microcrystals during catalysis. ICH exhibits a concerted helical displacement upon active-site cysteine modification that is gated by changes in hydrogen bond strength between the cysteine thiolate and the backbone amide of the highly strained Ile152 residue. These catalysis-activated motions permit water entry into the ICH active site for intermediate hydrolysis. Mutations at a Gly residue (Gly150) that modulate helical mobility reduce ICH catalytic turnover and alter its pre-steady-state kinetic behavior, establishing that helical mobility is important for ICH catalytic efficiency.
These results demonstrate that MISC can capture otherwise elusive aspects of enzyme mechanism and dynamics in microcrystalline samples, resolving long-standing questions about the connection between nonequilibrium protein motions and enzyme catalysis.
69
4.2. Introduction:
Protein dynamics play an important role in enzyme catalysis523–525,532. Many enzymes form covalent catalytic intermediates that can alter enzyme structure and conformational dynamics269,533–535. Because catalytic intermediates transiently modify enzymes, they may activate a new protein motion regime that facilitates later steps of catalysis. Determining how these transient changes in enzyme structure and dynamics facilitate passage along the reaction coordinate is a topic of intense interest in structural enzymology 525,535. Of all amino acids in proteins, reactive cysteine residues are subject to the most diverse set of covalent modifications348,536. Moreover, the targeted modification of cysteine residues proximal to active sites has become a frontline strategy for the development of covalent enzyme inactivators as tools for chemical biology and drugs537. Therefore, understanding how the H-bonding environment of active-site cysteine residues modulates their reactivity and how the covalent modification of cysteine thiolate anions controls protein conformational dynamics can provide general insights into the functions of many proteins whose activities require or are regulated by covalent modifications.
X-ray crystallography provides an atomically detailed ensemble and time- averaged view of protein conformational dynamics in the lattice environment538–541.
Technological advances in X-ray sources, detectors, and sample delivery have enabled a new class of crystallography experiments that report on nonequilibrium protein motions in response to external perturbations542. Because these perturbations can be selected for functional relevance (e.g., by the infusion of substrate or induction of a particular modification), the resulting nonequilibrium changes often correspond to functionally 70 important protein motions. In parallel, advances in computer modeling of multiple conformational substates now enable visualizing minor populations of crystalline protein structural ensembles. These approaches have permitted identification of networks of side- chain disorder543, allowed refinement of whole-protein ensemble models544, aided identification of minor binding modes of therapeutic ligands545, and characterized spatial correlations in protein mobility546.
The enzyme studied here is isocyanide hydratase (ICH; EC 4.2.1.103)
(PDB:3NON), a homo-dimeric cysteine-dependent enzyme in the DJ-1 superfamily that irreversibly hydrates a wide range of isocyanides to yield N-formamides32(Fig. 1A). Prior crystallographic studies of ICH showed evidence of helical mobility near the active-site cysteine101 residue (Fig. 1C), which coincided with photooxidation of C101 to sulfenic acid (C101-SOH)31. C101 accepts an S−-HN hydrogen bond from the severely strained
(φ = 14°, ψ = −83°) backbone amide of Ile152. When this hydrogen bond is disrupted in an engineered C101A mutant (Fig. 1C), Ile152 relaxes to an unstrained backbone conformation concomitant with displacement of the helix31,547.
The relevance of these motions for ICH function was not established in these prior studies. Here, we use X-ray crystallography, molecular dynamics simulation, mutagenesis, and enzyme kinetics to characterize a set of correlated conformational changes that remodel the ICH active site and propagate across the dimer interface in response to cysteine modification. Using time-resolved mix-and inject serial crystallography (MISC) at an X-ray free electron laser (XFEL), we directly observe formation of a covalent intermediate and transient, functionally important motions during
ICH catalysis. Pre-steady-state kinetics of wild-type and mutant ICH suggests that 71 cysteine-gated conformational changes are important for water to access and hydrolyze the catalytic intermediate. The prominence of conformational dynamics in ICH and the tractability of this protein using multiple biophysical techniques make ICH a valuable model system for understanding how cysteine modification can modulate functional protein dynamics.
4.3 Materials and Methods:
Crystallization, data collection, and processing: Pseudomonas fluorescens ICH enzyme was expressed from pET15b constructs in E. coli BL21DE3 cells as a thrombin- cleavable, N-terminally 6xHis-tagged protein and purified as previously described31. The
G150A and G150T mutants were made by site directed mutagenesis using mutagenic primers and were verified by DNA sequencing (Eurofins). All of the final proteins contain the vector-derived amino acids “GSH” at the N-terminus. Wild-type ICH, the
G150A, and the G150T mutants were crystallized by hanging drop vapor equilibration by mixing 2 μL of protein at 20 mg/ml and 2 μl of reservoir (23% PEG 3350, 100mM Tris-
HCl, pH 8.6, 200 mM magnesium chloride and 2 mM dithiotheritol (DTT)) and incubating at 22°C. Spontaneous crystals in space group P21 appear in 24-48 hours and were used to micro-seed other drops after 24 hours of equilibration. Micro-seeding was used because two different crystal forms of ICH crystals grow in the same drop and those in space group P21 are the better-diffracting crystal form. Notably, seeding G150T ICH using wild-type ICH crystals in space group P21 results exclusively in G150T crystals in space group C2 with one molecule in the asymmetric unit (ASU).
Cryogenic (100K) synchrotron data (PDB 3NON) were collected at the Advanced
Photon Source beamline 14BM-C from plate-shaped crystals measuring ~500x500x150 72
μm that were cryoprotected in 30% ethylene glycol, mounted in nylon loops, and cooled by immersion in liquid nitrogen as previously described31. Ambient temperature (274 K and 277 K) synchrotron data sets were collected at the Stanford Synchrotron Radiation
Lightsource from crystals mounted in 0.7 mm diameter, 10 μm wall thickness glass number 50 capillaries (Hampton Research) with a small volume (~5 μl) of the reservoir to maintain vapor equilibrium. Excess liquid was removed from the crystal by wicking, and the capillary was sealed with beeswax. For the 274 K dataset, the crystal was mounted with its shortest dimension roughly parallel to the capillary axis, resulting in the
X-ray beam shooting through the longest dimensions of the crystal during rotation. For the 277 K dataset, a large single capillary-mounted crystal was exposed to X-rays and then translated so that multiple fresh volumes of the crystal were irradiated during data collection. This strategy reduces radiation damage by distributing the dose over a larger volume of the crystal. This translation of the sample was factored into the absorbed dose calculation by assuming every fresh volume of the crystal received no prior dose, which is a best-case scenario.
The 100 K and 277 K datasets were collected on ADSC Q4 CCD detectors using the oscillation method. Separate low- and high-resolution passes were collected with different exposure times and detector distances and merged together in scaling, as the dynamic range of the diffraction data was larger than that of the detector. For the 274 K data, a Pilatus 6M pixel array detector (PAD) was used with shutterless data collection.
Because of the very high dynamic range of the detector, a full dataset was collected in a single pass ~5 minutes. The 274 K datasets were indexed and scaled using XDS548 while the 277K datasets were indexed and scaled and HKL2000549. The 100 K dataset was 73 processed as previously described 31. See Tables 2-3 for Crystallographic data refinement statistics.
Microcrystal growth and XFEL sample delivery. For the XFEL experiment, microcrystals of ICH (~10-20µm) were grown by seeding. Initial seeds were obtained by pulverizing macroscopic crystals via vortexing with 0.5 mm stainless steel balls for 5 minutes. A dilution of this seed stock was added to 31% PEG 3350, 250 mM MgCl2, 125 mM Tris-HCl pH=8.8 and then an equal volume of 40 mg/ml ICH was added and rapidly mixed. The dilution of the seed stock dictated the final size of the microcrystals and was optimized to produce crystals measuring ~20 μm x 20 μm x 2 μm. The mixture was incubated at room temperature with shaking for ~20 minutes until microcrystal growth stopped.
Samples were delivered to the beam using the concentric-flow microfluidic electrokinetic sample holder (coMESH) injector550 under atmospheric pressure conditions at room temperature and under normal atmosphere. The sample was loaded in a custom stainless steel sample reservoir. A Shimadzu LD20 HPLC pump hydraulically actuated a teflon plunger to advance the sample slurry. A stainless steel 20 µm frit filter (IDEX-HS) was placed after the reservoir in order to ensure smaller crystal sizes and mitigate clogging. The reservoir and filters were connected to the coMESH via a 100 µm x 160
µm x 1.5 m fused silica capillary (Molex). This capillary continued unobstructed through the center of the microfluidic tee (Labsmith) and terminated at 0 mm, 2 mm, or 200 mm recessed from the exit of a concentric 250 µm x 360 µm capillary, corresponding to Free
(Apo), 15 s and 5 min time delays, respectively. The capillaries were optically aligned to obtain a 0 offset, then were recessed and measured externally to achieve the 2 mm and 74
200 mm recess. In the case of the longer 200 mm offset, an additional tee (IDEX-HS) was added to increase the length. The outer line of the coMESH flowed the stabilizing solution (31% PEG 3350, 250 mM MgCl2, 125 mM Tris-HCl pH=8.8) in the case of the
Free (Apo) structure and flowed ~2 mM p-NPIC substrate in stabilizing solution for the
15 s and 5 min timepoints. DTT was omitted from all solutions as p-NPIC is not stable its presence and the p-NPIC substrate was at saturation (estimated concentration is ~2 mM) in this solution. The p-NPIC substrate was loaded into a 500 or 1000 µL gas tight syringe
(Hamilton). The stainless steel, blunt tip removable needle was connected to a 250 µm x
360 µm x 1 m fused silica capillary (Molex) and connected to the side of the coMESH microfluidic tee junction for the APO and 15 s delay and into the side of the Idex tee for the 5 minute delay. The syringe was driven by a syringe pump (KDS Legato 200) at 1-2
µl/min while being charged at 3.1 kV (Stanford Research Systems, SRS PS300) at the wetted stainless-steel needle.
The outer liquid focused the fluids ~500 µm away from the outer capillary towards the XFEL focus, of approximately 3 µm at the Macromolecular Femtosecond
Crystallography (MFX) end-station. The charged meniscus was approximately 5 mm away from a grounded counter electrode to complete the electro kinetic focusing. The time delays were assumed to be sufficiently long as compared to the electro kinetic mixing phenomena and were determined simply by the time the bulk fluid would traverse the offset distance. The flow rates and voltages were held constant during each time point and the flow of the crystals were optically monitored with a 50x objective to assure minimal flow deviations. For the APO (no delay) time point, the flow was 0.5 µl/min for both the sample and outer flows. For the 15 s time delay, the capillaries were recessed 2 75 mm, and the flows were 1 and 2 µl/min for the sample and p-NPIC flow rates, respectively. The combined bulk flows (1+2 = 3 µl/min) and the 2 mm offset dictated the delay time of 15 s. For the 5 min time delay, the capillaries were offset 200 mm and both flow rates were held at 1 µL/min, for a combined bulk flow of 2 µL/min. Overall, ~0.5 mL of sample crystals and 0.5 mL of 3 mM p-NPIC were consumed to gather this data.
XFEL Data collection and Processing. Data were collected at the MFX endstation of the Linac Coherent Light Source (LCLS). X-ray pulses were 9.5 keV, ~1x1012 photons per 40 fs pulse at 30 Hz repetition. Beam size at sample was ~3 μm. Diffraction data were collected on a Rayonix MX340-XFEL CCD detector operated in 4x4 binning mode and hits were analyzed in real time using OnDA 269. The XFEL diffraction data were processed with cctbx.xfel551, to identify 76,174 crystal hits for the Free (Apo) data, 53,476 crystal hits for the 15 second delay data and 36,875 crystal hits for the 5 minute delay data. A small fraction of multiple lattice hits (<0.01% of total) were shown to have little impact on data quality when included in the final post-refinement steps. For the Free (Apo) data, 23,669 frames were indexed and 21,834 of those had their intensities merged and integrated. For the 15s data, 26,507 frames were indexed and 24,286 of those had their intensities merged and integrated. For the 5min data, 16,893 frames were indexed and 14,483 of those had their intensities merged and integrated. To correct the intensity measurements and merge the data, postrefinement was carried out with three cycles of PRIME552 using model unit cell dimensions from a synchrotron wild type ICH structure31. The Lorentzian partiality model used parameters gamma_e = 0.001, frame_accept_min_cc = 0.60, uc_tolerance = 5, sigma_min = 2.5, and partiality_min =
0.2. The X-ray crystallographic data statistics for all datasets are provided in Tables 2-3. 76
Selection of mix-and-inject X-ray time delays. We collected absorbance spectra of ICH crystals in reservoir solution supplemented with 1mM p-NPIC and 40% glycerol using
UV-vis microspectrophotometry at SSRL BL 11-1 (Fig. 1B). We freeze-trapped the reaction in the cryostream at fixed time-delays after adding substrate to the crystal before collecting a spectrum. The p-NPIC absorption maximum is ~270 nm, while p-nitrophenyl formamide product absorption maximum is ~315 nm on this instrument. The lack of an isosbestic point suggests an intermediate accumulated. At a time delay of 15s, the substrate peak at 270 nm had disappeared, while the product peak at 315 nm had not formed yet. At 5 minutes, substrate is exhausted while the product peak at 315 nm had fully formed, informing the timing for the XFEL experiment. The macroscopic (~2x106
μm3) crystals from which these spectra were collected were damaged by exposure to substrate and did not diffract X-rays well after substrate was introduced, precluding a cryotrapping X-ray diffraction experiment. In contrast, the microcrystals used for the
XFEL experiment (~2x103 μm3) were not damaged by substrate, necessitating a serial X- ray crystallography approach.
Crystallographic model refinement. All refinements were performed in PHENIX1.9 against structure factor intensities using individual anisotropic ADPs and riding hydrogen atoms553. Weights for the ADP refinement and relative weights for the X-ray and geometry term were optimized. After convergence of the isotropic ADP refinements, translation-libration-screw (TLS) refinements554 with automatic rigid body partitioning were performed in PHENIX. For the C101-SOH and G150A ICH synchrotron datasets, the alternate conformation for helix H (residues 148-173 in chain A) was modeled as a single occupancy group, reflecting the presumed correlated displacement of sidechain 77 and backbone atoms. Other disordered regions were also built into multiple
555 conformations manually in COOT if supported by 2mFo-DFc and mFo-DFc electron density maps. Occupancies of these groups were refined but constrained to sum to unity in PHENIX. Omit maps were calculated after removing the areas of interest from the model and refining for three macrocycles. Final models were validated using
MolProbity556 and the validation tools in COOT and PHENIX. Model statistics are provided in Table 3. Isomorphous difference maps (Fo(15s) – Fo(XFELAPO) and Fo(SR) –
Fo(XFELAPO)) were calculated using data to the lowest high resolution of the data sets,
1.55 Å. Structure factor data and refined coordinates are available in the Protein Data
Bank with the following accession codes: 6NI4, 6NI5, 6NI6, 6NI7, 6NI8, 6NI9, 6NJA,
6NPQ, 6UND, and 6UNF.
Evidence of ICH in-crystallo catalysis.. A single crystal of ICH was mixed with 1 mM substrate, para nitrophenyl isocyanide (p-NPIC), incubated for the indicated times in seconds (Fig. 1B), mounted and cryocooled to 100 K in a nitrogen cryo-stream.
Absorbance spectra were collected using the inline spectrophotometer at BL 11-1 at the
Stanford Synchrotron Radiation Lightsource (SSRL). pH gradient wildtype ICH crystal growth, data collection and processing. For the pH screen, crystallization buffers were made with 23% PEG 3350, 0.2 M MgCl2, 2mM DTT and 0.1M Tris-HCl or 0.1M Na Acetate or 0.1M Na Citrate. Wildtype ICH was crystallized by hanging drop vapor equilibration by mixing 2 μL of protein at 20 mg/ml and 2 μl of the corresponding reservoir solution and incubating at 22°C. The drops were streak seeded to facilitate the growth of the the better diffracting P21 space group. 78
Synthesis of para-nitrophenyl isocyanide (p-NPIC). ICH will accept diverse
32 isocyanide substrates . Due to its strong absorption in the visible range (λmax=320 nm), para-nitrophenyl isocyanide (p-NPIC) was used here. The p-NPIC substrate is not commercially available and was synthesized by the Dr. David Berkowitz and his group
(University of Nebraska Lincoln, Department of Chemistry) via dehydration of the N- formamide precursor. To synthesize N-(4-Nitrophenyl) formamide, a mixture of amine
(10 mmol) and formic acid (12 mmol) was stirred at 60 °C for 2h.
After completion of the reaction, as judged by Thin Layer Chromatography (TLC), the mixture was diluted with CH2Cl2 (100 mL), washed with saturated solution of NaHCO3 and brine (100 mL), and then dried over anhydrous Na2SO4 and concentrated under reduced pressure. The residue was dissolved in hot acetone and crystallized, to yield N-
(4-Nitrophenyl) formamide (1.58 g, 96%) as a yellow solid. The product was verified using 1H NMR, 13C NMR, IR, and Time of Flight (TOF) mass spectra.
To synthesize p-Nitrophenyl isocyanide, the mentioned N-(4-Nitrophenyl) formamide product (8.0 mmol) and triethylamine (56 mmol) were dissolved in
CH2Cl2 (30 mL) and cooled to -10 °C.
79
A solution of triphosgene (8.0 mmol) in CH2Cl2 (50 mL) was slowly added and stirring was continued at -10 °C for another 30 minutes. After completion of reaction (judged by
TLC), the reaction mixture was passed through a pad of silica gel and washed with
CH2Cl2. The filtrate was dried over anhydrous Na2SO4, and then concentrated under reduced pressure. The residue was triturated with cold hexane to yield para-nitrophenyl isocyanide (1.36 g, 92%) as a yellow solid. 1H NMR, 13C NMR, IR, and Time of Flight
(TOF) mass spectra were collected to verify the product structure. The para-nitrophenyl isocyanide product was stored as a dry solid at -80°C until needed.
ICH enzyme kinetics. Steady state ICH rate measurements were initiated by the addition of ICH (final concentration of 1 µM) to freshly prepared p-NPIC solutions ranging from
0-60 µM in reaction buffer (100 mM KPO4 pH=7.0, 50 mM KCl, and 20% DMSO). p-
NPIC was diluted from a 0.5 M stock solution in dimethyl sulfoxide (DMSO), which was stored at -80 °C and protected from light. A 200 µl reaction was maintained at 25 °C in a
Peltier-thermostatted cuvette holder. The formation of the product, para-nitrophenyl formamide (p-NPF) was monitored at its absorption maximum of 320 nm for two minutes using a UV-Vis Cary 50 Spectrophotometer (Varian, Palo Alto, CA). A linear increase in A320 was verified and used to calculate initial velocities. Isocyanides can spontaneously hydrolyze slowly in aqueous solutions, with the rate increasing as pH is lowered. To ensure that all measured product formation was due to ICH catalysis, the rate of spontaneous p-NPIC hydrolysis was measured without added ICH. These values were near the noise level of the spectrophotometer and were subtracted from the raw rate measurements before fitting the Michaelis-Menten model. The extinction coefficient at 80
320 nm for p-NPF was determined by using ICH to convert known concentrations of p-
NPIC to p-NPF, followed by measuring the absorbance at 320 nm. The slope of the resulting standard curve was defined as the extinction coefficient of p-NPF at 320 nm;
4 -1 -1 ε320=1.33x10 M cm . This p-NPF ε320 value was used to convert the measured rates from A320/sec to [p-NPF]/sec. All data were measured in triplicate or greater and mean values and standard deviations were plotted and fitted using the Michaelis-Menten model as implemented in Prism (GraphPad Software, San Diego, CA). Reported Km and kcat values and their associated errors are given in Fig. 4E-F.
Pre-steady state ICH kinetics were measured at 25 ˚C using a Hi-Tech KinetAsyst stopped flow device (TgK Scientific, Bradford-on-Avon, United Kingdom). Data were collected for each sample for two seconds (instrument deadtime is 20 ms) in triplicate.
The final enzyme concentration after mixing was 10 M and final p-NPIC concentrations were 20, 40, 80, 120, 160, and 320 M. Product evolution was monitored at 320 nm using a photodiode array detector. Kinetic Studio software (TgK Scientific, United
Kingdom) was used to analyze the kinetic data and to fit a mixed model containing a single exponential burst with a linear steady state component: -Aexp(-kt) + mt + C. In this equation, t is time, k is the burst phase rate constant, m is the linear phase (steady- state) rate, A is the amplitude of the burst component, and C is a baseline offset constant.
The linear slope m was used to calculate steady state turnover numbers, which agree well with kobs values obtained from steady state kinetic measurements at comparable substrate concentrations. Single turnover experiments were also performed using final concentrations of 40 M enzyme and 20 M p-NPIC after manual mixing. Spectra were collected using a Cary 50 spectrophotometer (Varian, Palo Alto, CA, USA). Spectra 81 were collected every 10 seconds with a deadtime of ~30 seconds after manual addition of enzyme.
Isocyanide (p-NPIC) toxicity assays. E coli BL21DE3 cells bearing the ICH containing pET15b recombinant plasmids were grown until OD600=0.5-0.6 at 37˚C and 250rpm shaking, upon which they were induced with 0.5mM IPTG. 1 ml of induced cells were mixed rapidly with 5mls of cooled autoclaved soft agar (0.8%) supplemented with
100mg/ml Ampicillin and poured over 1.5% LB agar plates and incubated at room temperature (22-25˚C) for 1 hour. Once the soft agar had solidified, stress chemicals were prepared fresh (500mM, 100mM and 10mM p-NPIC in 100%DMSO). Each LBA plate was divided into four quadrants and sterile autoclaved filter discs (1 cm diameter) were placed in the four sections. 10µl stress chemical was added to each disk and the plates were incubated at 37˚C overnight. Diameters of the developed zone of clearances were manually measured in triplicates after 24 hours and plotted as a function of substrate concentration using Prism (GraphPad Software, San Diego, CA).
4.4 Results:
4.4.1 ICH actively catalyzes hydration of isocyanides to formamides in crystallo.
Prior to XFEL characterization of catalytic turnover of ICH, it is important to ensure that the enzyme is active in the crystal form as a pre-requisite to the planned
XFEL MISC experiment. A single crystal of ICH was mixed with 1 mM p-NPIC, mounted and cryocooled to 100 K in a nitrogen cryostream. Absorbance spectra were collected using the inline spectrophotometer at BL 11-1 at the Stanford Synchrotron
Radiation Lightsource (SSRL). On this instrument, the p-NPIC substrate has an absorption maximum of 270 nm and the p-NPF product has an absorption maximum of 82
315 nm. Note the absence of an isosbestic point (Fig. 1B).
4.4.2 Formation of thioimidate reaction intermediate during ICH catalysis triggers active site conformational dynamics in ICH.
Mix and Inject Serial Crystallography (MICS) at the LCLS XFEL at 298K allowed us to examine the role of active site structural dynamics in ICH turnover by directly monitoring catalysis in ICH microcrystals. We infused the substrate p-nitrophenyl isocyanide (p-NPIC) into a stream of unmodified ICH microcrystals (Methods) and collected diffraction data at carefully selected time delays (Fig. 1B). The concentration of p-NPIC in the stream allowed only a single turnover of crystalline ICH before substrate depletion. It is important to note here that due to the “diffraction before destruction” principal, SFX data are minimally affected by X-ray radiation damage557, as a result of which the enzyme suffered no radiation-induced oxidation of C101 (Fig. 1D). It is essential to rule out radiation damage induced conformational switching in ICH from catalysis induced dynamics because both events are triggered by C101 covalent 83
Fig 1:(A) Hydration of isocyanides by ICH yields corresponding formamides. (B) In-crystallo ICH turnover monitored in a single crystal mixed with 1 mM p-NPIC, mounted, cryocooled to 100 K in a nitrogen cryostream. Absorbance spectra collected using the inline spectrophotometer at BL 11-1 at the Stanford Synchrotron Radiation Lightsource (SSRL). (C) The ICH dimer is shown as an overlay of WT ICH (purple) and C101A ICH (blue). The “B” protomer is shown in yellow. The mobile helix H in C101A and areas exhibiting correlated backbone–side-chain disorder are rendered opaque. Ile152 is a Ramachandran outlier (Inset) whose backbone torsion angles move with helical displacement.(D) ICH completes a full catalytic cycle in the crystal during MISC. Helix H is shown with 2mFo-DFc electron density (0.8 rmsd) prior to the introduction of substrate (blue) and after substrate has been exhausted (orange). These maps overlap almost perfectly, indicating that ICH is fully restored to its resting conformation after catalysis. The hydrogen bond between the peptide backbone of Ile152 and Cys101 is shown in a dotted line. The helix is not mobile in these resting structures, indicated by the absence of features in the mFo-DFc difference electron density (2.5 rmsd prior to substrate [green] and after catalysis is complete [purple]) (E) MISC confirms that ICH forms a covalent Cys101-thioimidate intermediate 15 s after substrate mixing. Difference mFo-DFc electron density contoured at 2.5 rmsd (green) supports sampling of shifted helix H conformations upon intermediate formation. (F) Postulated reaction mechanism for ICH, beginning with Cys101 thiolate attack at the electrophilic carbenic carbon atom of isocyanide substrates and proceeding in the direction of the arrows. A previously postulated thioimidate intermediate (red box) eliminates the charge on Cys101 and weakens the H-bond to Ile152 (dashed red line). This relieves backbone torsional strain at Ile152 and permits sampling of shifted helix H conformations (green curved arrow), allowing water access to the intermediate for hydrolysis. modification. The proposed reaction mechanism of ICH (Fig. 1F). states that the
catalytic C101 nucleophile attacks organic isocyanides at the electron-deficient carbene- 84 like center, followed by proton abstraction from a nearby general acid to generate a covalent thioimidate intermediate. An enzyme-linked thioimidate, which has not been previously observed, eliminates the negative charge of the C101 thiolate (S−) and is proposed to reduce the strength of the C101-I152 hydrogen bond.
Fig 2: Observation of thioimidate intermediate formation and enhancement of ICH helix mobility. In (A), omit mFo-DFc electron density is shown at 3.0 rmsd (green) and 5.0 rmsd (purple), unambiguously demonstrating presence of the catalytic intermediate. In (B-D), anisotropic atomic displacement parameters (ADPs) from TLS models refined against the XFEL datasets are shown at the 75% probability level and colored according to Beq value, from blue (7 Å2) to red (30 Å2). Starting with the resting enzyme before substrate is introduced (B), helical mobility is transiently elevated upon thioimidate intermediate formation during catalysis (C) and then is reduced again upon completion of catalysis (D) once substrate is consumed 5 minutes after mixing.
After 15 s of mixing with substrate, 2mFo-DFc (Fig. 1E) and omit mFo-DFc electron density maps (Fig. 2A) unambiguously reveal the formation of a thioimidate covalent intermediate in both ICH protomers. Once the intermediate is formed, positive mFo-DFc difference electron density appears around helix H (Fig. 1E) and its B-factors increase
(Fig. 2B-D), indicating that it samples a new conformational ensemble. This difference density and elevated B-factors are absent both prior to catalysis and after ICH has 85
exhausted substrate 5 minutes after mixing (Fig. 1D, Fig. 2B-D). Therefore, helix H
becomes transiently mobile upon intermediate formation during ICH catalysis, changing
the active site microenvironment.
Fig 3: X-ray induced cysteine oxidation drives helical motion in ICH. (A-C) The top panels show the environment of Cys101 with varying degrees of oxidation to Cys101-sulfenic acid. 2mFo-DFc electron density is contoured at 0.7 RMSD (blue) and the hydrogen bond between the peptide backbone of Ile152 and Cys101 is shown in a dotted line. “Cryo” is synchrotron data collected at 100 K (PDB 3NON), “RT less oxidized” is synchrotron data collected at 274 K with an absorbed dose of 2.4x104 Gy, and “RT more oxidized” is synchrotron data collected at 277 K with an absorbed dose of 3.7x105 Gy. The lower panels show helix H in its strained (black) and relaxed, shifted conformations (grey). 2mFo- DFc electron density is contoured at 0.8 RMSD (blue) and omit mFo-DFc electron density for the shifted helical conformation is contoured at 3.0 RMSD (green). At 274-277K, increased Cys101 oxidation disrupts the hydrogen bond to Ile152 and results in stronger difference electron density for the shifted helix conformation. (D) The refined occupancy of helix H for each X-ray dataset, indicating that increases in temperature and Cys101 oxidation result in higher occupancy for the shifted (relaxed) helix conformation. (E) Mechanism of X-ray induced covalent modification of C101 and weakening of the S— -NH hydrogen bond. (F) Cys101 oxidation leads to a weakening of the Ile152-Cys101 H-bond. Electrostatic Poisson-Boltzmann surfaces (red negative, blue positive charge) calculated from the Cys101-Ile152 (A) and Cys-SOH-Ile152 (B) environments. Cysteine photooxidation neutralizes the negative charge of the sulfur atom, weakening the N-H ..S hydrogen bond. We calculated a reduction in the Cys101-Ile152 hydrogen bond energy from -2.2 kcal/mol under reduced conditions (i.e. with a thiolate acceptor) to -0.91 kcal/mol upon Cys101-SOH formation. 86
We reasoned that the on-pathway thioimidate intermediate weakens the interactions that hold helix H in a strained conformation similarly to the X-ray-induced C101-SOH modification in the resting enzyme. Consistent with this idea, the cryogenic dataset collected at 100K (Cryo) also shows evidence of radiation-driven C101-SOH formation and a corresponding increase in helical mobility31 (Fig. 3A). To enrich populations of the conformational shifts, we therefore collected a series of X-ray diffraction data sets of WT
ICH at increasing doses of X-ray radiation (Fig. 3A-C). Two ambient temperature (274-
277K) synchrotron radiation datasets at 1.20-1.15 Å reveal a radiation-dose dependent
C101 oxidation and helical displacement with increasing occupancy (Fig. 3D). Refined occupancies of alternate conformations of helix H confirm a population shift towards the displaced helix position along this series of data sets that correlates with the extent of absorbed X-ray dose and concomitant C101 oxidation. The shifted helix conformations across data sets are highly similar, suggesting that the observed electron density features result from coupling between this helix and specific, radiation-induced photochemistry at
C101.
Notably, these stronger density peaks correspond to the difference electron density observed in the MISC experiment during ICH catalysis (Fig. 1E, Fig. 3A-C). In the ambient temperature synchrotron radiation datasets, the helical shift is caused by radiation-induced oxidation of the C101 eliminating the negative charge on Sγ thiolate and thus weakening the hydrogen bond between the amide H of Ile152 and C101 Sγ, from -2.2 kcal/mol with a thiolate acceptor to -0.91 kcal/mol with a C101-SOH acceptor
(Fig. 3E-F). Similar loss of C101 thiolate negative charge occurs in the on-pathway thioimidate intermediate, which also weaken the C101-I152 hydrogen bond. (Fig. 1F). 87
4.4.3 Cysteine modification-gated motions increase ICH catalytic efficiency
To modulate helical mobility and measure the impact on ICH catalysis, we designed two mutations, G150A and G150T. G150 is part of a highly conserved diglycyl motif that moves ~3Å to accommodate helical motion in ICH. Ambient temperature
(274-277 K) crystal structures of G150A and G150T ICH show that the added steric bulk at this position biases the helix towards its relaxed conformation (Fig. 4A, B). In G150A
ICH, helix H samples both conformations in both protomers (Fig. 4A, C-D). By contrast, the G150T structure shows helix H constitutively shifted to its relaxed position (Fig. 4B).
Both G150A and G150T have an alternate C101 sidechain conformation which conflicts with the unshifted conformation of Ile152 and helix H (Fig. 4A-B asterisk), indicating that the helix must move partially independently of C101 modification in these mutants.
As in wild-type ICH, G150A shows evidence of C101-SOH oxidation in the electron density (Fig. 4A). In contrast, C101 in G150T ICH is not modified by comparable exposure to X-rays (Fig. 4B), indicating that the Ile152-NH-C101-S- hydrogen bond is important for enhancing C101 reactivity. 88
Fig 4: (A-B) (Top). 2mFoDFc electron density is contoured at 0.7 rmsd (blue) and the H-bond appears as a dotted line. Both G150 mutations permit unmodified Cys101 to sample conformations (asterisk) that sterically conflict with Ile152 in the strained helical conformation (black), requiring the helix to sample shifted conformations (gray) in the absence of Cys101 modification. (A and B, Lower) The helix in its strained (black) and relaxed, shifted conformations (gray). 2mFo-DFc electron density is contoured at 0.8 rmsd (blue) and omit mFo-DFc electron density is contoured at +3.0 rmsd (green).(C-D) Helical mobility is asymmetric in wild-type but not G150A ICH The top panels show the environment of Cys101 in the second protomer (“protomer B”) of wild-type (WT) and G150A ICH. 2mFo-DFc electron density is contoured at 0.7 RMSD (blue) and the hydrogen bond between the peptide backbone of Ile152 and Cys101 is shown in a dotted line. The lower panels show the helix in its strained (black) and relaxed, shifted conformations (grey). 2mFo-DFc electron density is contoured at 0.8 RMSD (blue) and omit mFo-DFc electron density for the shifted helical conformation is contoured at 3.0 RMSD (green). Wild-type ICH does not show strong evidence of a second conformation, even though Cys101 has been partially photooxidized to Cys101-SOH (top). In contrast, the helix in monomer B of G150A ICH shows strong evidence of a second, relaxed conformation, similar to monomer A. Steady-state (E) and pre-steady state (F) enzyme kinetics of wild-type (WT; blue circles), G105A (black squares), and G150T (red triangles) ICH. Both the G150A and G150T mutations result in similar decreases in steady-state kinetics compared to WT enzyme. (E) The burst rate constant in pre-steady state kinetics is linearly dependent on substrate concentration. This linear dependence indicates a second order rate process during the burst phase, consistent with thioimidate intermediate formation. WT and G150A have similar second order burst rate constants (G) Pre-steady-state enzyme kinetics of WT (blue circles), G105A (black squares), and G150T (red triangles) ICH at 160 μM p-NPIC shows a pronounced burst phase for each protein with differing burst and steady-state rate constants. (H) Single-turnover spectra of ICH enzymes with p-NPIC substrate shown from early (red) to later (blue) timepoints in 5-s increments. At early times, G105A and G150T accumulate a species with λmax = 335 nm, likely the thioimidate intermediate that resolves to product in the blue spectra with λmax = 320 nm. (I) The catalytic cycle of ICH (Summary figure). 89
Unlike the C101A mutant, which shows similar helical displacement but is catalytically inactive, G150A and G150T ICH are catalytically active. This allowed us to investigate the role of helical displacement in the ICH catalytic cycle. Steady-state enzyme kinetics of the G150A and G150T mutants measured using p-NPIC as the substrate show a ~6–fold reduction in kcat for both mutants compared to wild-type enzyme but largely unchanged KM values (Fig. 4E, F Table 1). In contrast to their similar steady-state kinetic behavior, the G150 mutants have divergent pre-steady state kinetic profiles in stopped-flow mixing. ICH exhibits “burst” kinetics (Fig. 4G), indicating that the rate-limiting step for ICH catalysis comes after formation of the thioimidate intermediate. G150A ICH has a burst exponential rate constant k of ~11 s-1 that is comparable to the wild-type enzyme, but G150T has a reduced burst rate constant of ~4 s-
1, indicating a slower chemical step (Fig. 4E-G, Table 1). WT and G150A ICH have comparable second order rate constants for the burst phase, while G150T is markedly lower (Fig. 4E-F, Table 1). In addition, the burst amplitude of G150A is approximately twice as large as that of the wild-type ICH despite equal concentrations of enzyme (Fig.
4G), which correlates with both protomers of G150A having a mobile helix H compared to only one protomer of wild-type ICH (Fig. 4C-D). This suggests communication between the two active sites in the ICH dimer that is dynamically gated by helix H.
Although both the G150A and G150T mutations impair ICH catalysis, the kinetic effect of the G150A mutation is predominantly in steps after formation of the intermediate, while G150T impairs both the rate of intermediate formation and later, rate- limiting steps. Thus, modulation of helical dynamics by the G150A and G150T mutants have divergent effects on ICH catalysis, indicating that the active site helical switching is 90 essential to support efficient ICH catalysis. Furthermore, we observe spectral evidence for the thioimidate intermediate in single-turnover UV-visible spectra (Fig. 4H). G150A
ICH accumulates a species whose absorbance maximum is 335 nm, while WT ICH completes a single turnover and accumulates the 320 nm formamide product in the ~30 second deadtime of manual mixing (Fig. 4H). G150T ICH accumulates less of the 335 nm intermediate than G150A ICH due to a closer match between the rates of formation and consumption of the intermediate in G150T ICH. In G150A ICH, the 335 nm species slowly converts to the 320 nm product over ~40 s, consistent with the slow rate of product formation observed after the burst in G150A pre-steady state kinetics (Fig. 4G).
These data support the conclusion that the 335 nm species is the ICH-thioimidate intermediate observed in the MISC experiment and that G150A is impaired in hydrolyzing this covalent intermediate from the active site C101 due to perturbed dynamics of helix H. It will be interesting to use the lower G150A mutant ICH in a similar MISC experiment at LCLS at ambient temperature to potentially trap the other reaction intermediate (tetrahedral) during IC
The impaired catalytic effects of the G150A and T mutations can also be demonstrated in in vivo p-NPIC toxicity assays performed with E.coli BL21DE3 cells bearing the pET15b constructs (Fig. 5A-D). BL21DE3 cells expressing the catalytically dead enzyme C101S was used as negative 91
Fig. 5: (A-C) LBA plates showing zones of clearances in wtICH, G150T and G150A respectively, in response to varying doses of isocyanide p-NPIC (DMSO vehicle control, 10mM, 100Mm, 500mM p- NPIC). (D) Diameters of the zones of clearances formed by E. coli BL21DE3 cells bearing the various ICH-pET15b constructs linearly track the concentration of the stress chemicals control and pET15b only cells were used as the empty vector controls. DMSO vehicle controls were included in each run (n=3). Fig. 5 clearly demonstrates the loss of isocyanide (p-NPIC) remediation abilities in BL21DE3 cells expressing G150A and
G150T (Fig. 5B-C) compared to wildtype ICH bearing cells (Fig. 5A) as seen by the increased diameters of their zones of clearances (Fig. 5D).
4.4.4 Propagation of cysteine-gated conformational changes across the ICH dimer.
To characterize the extent of the conformational response of the entire ICH dimer to the 92
Fig. 6: (A) Isomorphous difference maps show widely distributed conformational changes upon cysteine modification. Fo(SR) – Fo(XFELFree) (top panels) and Fo(XFEL15s) – Fo(XFELFree) (bottom panels) difference maps are contoured at 2.75 rmsd. difference features (green, positive; red, negative) are nonuniformly distributed in both maps, radiating out from the site of the catalytic nucleophile (yellow spheres). This is more pronounced in the A protomer (slate blue) than the B protomer (grey) in the Fo(SR) – Fo(XFELFree) map (top panels). Difference features generally show the same, but weaker pattern in the Fo(XFEL15s) –Fo(XFELFree) map, where the 15 s timepoint corresponds to thioimidate intermediate formation. The maps are phased with the Free (no substrate) XFEL structure. (B) Isomorphous Fo(SR) – Fo(XFELFREE) difference map reveals broadly altered structure and dynamics upon formation of the Cys101-SOH in the 274-K synchrotron radiation (FoRT) dataset. Helices H and I are opaque in both conformers. The catalytic nucleophile is shown in spheres. Difference electron density features are nonuniformly distributed, with stronger features near helix H in the A protomer, and along region B169–B189, which contacts the N-terminal end of helix H. Maps are contoured at ±3.0 rmsd. (C) CONTACT analysis identifies allosteric coupling across the dimer interface, in striking agreement with isomorphous difference maps and rmsfs from (B). The A protomer is color-coded in blue; the B protomer, in red. Residues identified in the CONTACT analysis are projected onto the cartoon. modification of C101, we calculated an Fo(SR) – Fo(XFELFree) isomorphous difference map, phased with a structural model obtained from the XFEL data set. The Fo(SR) –
Fo(XFELFree) isomorphous difference map provides an unbiased view of differences in molecular conformation between two data sets, reporting specifically on changes in ICH that occur in response to oxidation of C101. A Fo(15s) – Fo(XFELFree) map calculated using the thioimidate dataset after 15 s of mixing and the dataset prior to introduction of substrate (Free) reveals similar features to those seen in the Fo(SR) – Fo(XFELFree) map 93
(Fig.6A) signifying that the widespread dynamic response observed in Fo(SR) –
Fo(XFELFree) difference map is common to different types of C101 modification. We further examined the dynamical communication across the dimer interface in ICH using
CONTACT network analysis of the electron density maps. CONTACT elucidates pathways of collective amino acid main- and sidechain displacements through mapping van der Waals conflicts that would result from sidechain conformational disorder if correlated motions are not considered 543. CONTACT identified a large network of correlated residues in protomer A (with the mobile helix) that connects with a smaller network in protomer B (Fig. 6C), corroborating the isomorphous difference map. The key residues in CONTACT analysis that bridge the dimer interface are Tyr181 and Thr153, which are also key residues identified in the isomorphous difference map. Our results indicate that the conformational changes upon C101-I152 H-bond modification in the synchrotron structure correspond to later steps in the catalytic cycle, allowing helical motion that facilitates intermediate hydrolysis and product release. At the same time, increased allosteric transmission during these later steps may prime the dimer for the next catalytic cycle. Ongoing work includes generation of Y181F and D157N mutants to eliminate or weaken an intra-dimer hydrogen bonding between Y181 and D157 residues to manipulate this dimer spanning allosteric network of corelated motions to better elucidate its impact on ICH catalysis.
4.4.5 The charge on C101 in ICH modulates active site dynamics
pH can be used as a controlling parameter to modulate C101 gated active site helical dynamics in ICH, independent of X-ray induced oxidation or catalysis. The rationale behind this is, when the pH of the reservoir solution is lesser than the pKa of the 94
C101, it is protonated. Therefore, the protonated (neutrally charged) C101cannot effectively form the negative charge assisted H bond with Ile125, causing it to relax and adopt an outlier conformation, triggering helical mobility. In this study, wildtype ICH was crystallized at pH 4.2, 5.0, 5.4, 6.0 and 8.3 (Fig. 7). Upon optimization of pH in the crystallization buffer, ICH shows varying degrees of C101 protonation/deprotonation and corresponding helical mobility in both protomers.
At pH 4.2 and 5.0 (Fig. 7A-B), C101 is protonated and the Ile152 primarily adopts a relaxed conformation, in contrast to pH 6.0 and 8.3 (Fig. 7D-E), where the C101 is deprotonated (negatively charged); the negative thiolate forms hydrogen bond with
Ile152, holding it in the restrained conformation. It is to be noted here that the pH 5.6 and
8.3 datasets show some positive density around the C101, indicating some oxidation of the C101, which potentially contributes to the Ile152 relaxed conformation (negligible). It would be beneficial to repeat these datasets, with properly handled crystal samples to ensure there is no oxidation. pH 5.0 (Fig. 7C), shows a mixture of both protonated and deprotonated C101 and a disordered active site with Ile152 occupying both restrained and relaxed conformations. These studies conclude that when the C101 is negatively charged, the negative thiolate can form H bond with Ile152 and hold it in the restrained configuration, whereas when the C101 is neutral or uncharged, it cannot form H bond with Ile152, releasing it to its relaxed configuration, triggering active site helical dynamics. This work is important because it confirms that the helical dynamics in ICH are dependent on the overall charge on the C101. It also shows that pH can be used as a control parameter to tune the internal ICH active site motions, thereby making this 95 enzyme an excellent model for the development of Diffuse Scatter (DS) data analysis and processing methods.
Figure 7: pH can be used to control ICH helical motions (A) pH 4.2 and (B) pH 5.0 shows protonated (uncharged) Cys101 and Ile152 shifted to relaxed configuration. (C) pH 5.0 shows a mixture of protonated and deprotonated Cys101 and sampling of relaxed and restrained configurations by Ile152. (D) pH 6.0 and (E) pH 8.3 show deprotonated Cys101 and Ile152 primarily occupying restrained configuration.
Diffuse scattering occurs from an imperfect or disordered crystal, when the scattering amplitudes do not add up constructively as in Bragg diffraction. The amplitude of the
Bragg peak decreases and this lost intensity is redistributed into diffuse scattering. Thus, 96
DS has the potential to inform on corelated motions inside crystals, but technological challenges prevent use of DS in protein crystallography. Currently there exist limited tools to interpret and process DS data, as a result of which, the DS portion of datasets are most often ignored when refining models. By manipulating the internal motions in ICH, we can develop DS data analysis and processing methods.
4.5 Discussions:
Covalent modification is a common and physiologically important perturbation to proteins. Reactive residues such as cysteine are prone to diverse covalent modifications with catalytic or regulatory consequences. We found that cysteine modification in ICH remodels active site H-bonding networks and gates catalytically important changes in protein dynamics with a clearly defined mechanism. ICH catalyzes a reaction that can be divided into an early phase dominated by nucleophilic attack of the reactive C101 at the electrophilic carbenoid carbon of its isocyanide substrate and a later phase dominated by water attack at the thioimidate and weaker nucleophilicity of C101 (i.e. a better leaving group) to release the N-formamide product (Fig. 1F). Modification of the active site cysteine thiolate upon formation of the thioimidate, during the first step of catalysis, neutralizes its negative charge and weakens a key H-bond, activating correlated motions that span the entire ICH dimer. After formation of the thioimidate, the helix samples the shifted conformation due to weakening of the I152-C101 H-bond, dynamically remodeling the ICH active site (Fig. 1F, Fig. 4I) and permitting water entry. This transient remodeling is required for hydrolysis of the thioimidate intermediate, because the active site containing the intermediate in occluded and does not contain room enough for a water to enter without motion. Water attack at the C-S bond of the thioimidate is 97 proposed to form a tetrahedral intermediate (Fig. 1F).At this point, the H-bond between
C101 and I152 that defines the strained conformation of helix H can reform, thereby stabilizing the nascent C101 thiolate and making it a better leaving group, thereby coupling dynamical changes in the enzyme to progress along the reaction coordinate (Fig.
4I).With the strained helical conformation restored, the product is released and leaves the active site poised for another cycle of catalysis (Fig. 4I).
The divergent pre-steady state kinetics of the G150A and G150T mutants suggest a model where the strained helical conformation of ICH has the highest competence for the initial isocyanide attack by C101, forming the thioimidate intermediate (Fig. 1E, Fig. 2A). This is also consistent with the diminished propensity of
C101 for photooxidation in G150T, where the helix is constitutively shifted, suggesting that C101 is less reactive in this environment. The G150A/T mutants modulate helical dynamics and thus allow us to determine if the non-equilibrium helical motions that we observe in the thioimidate intermediate XFEL structure are incidental to or important for catalysis. Synchrotron X-ray crystallographic data indicate that the G150A mutation enhances sampling of shifted helical conformations even in the absence of C101 modification, and this shifted conformation is further populated once the C101-Ile152 H- bond is weakened by C101-SOH formation. In addition, G150A ICH accumulates a spectrally distinct 335 nm species that likely corresponds to the thioimidate intermediate detected by XFEL crystallography. Our interpretation of these data is that the thioimidate intermediate accumulates in G150A ICH owing to an impaired resetting of the strained helical conformation, which reduces the rate of thioimidate hydrolysis and enzyme turnover. Consistent with the pre-steady state kinetic data, this kinetic model for G150A 98
ICH predicts that the early chemical steps promoted by the strained conformation of helix
H would not be impaired by the mutation but that the steady-state rate would be significantly diminished. In contrast, the G150T mutation causes a constitutively shifted helix, reducing rates of both initial C101 attack at the isocyanide carbon atom in the first chemical step and thioimidate hydrolysis in later steps, as evidenced by the lower burst and steady state rates of G150T ICH. Therefore, the G150A and G150T mutations have divergent effects on the early steps of ICH catalysis but similar detrimental effects on the later, rate-limiting steps.
Protein structures fluctuate owing to thermal motion and in response to functional changes such as ligand binding. As a consequence, it is challenging to determine which protein motions are functionally most important at equilibrium. Enzymes that are transiently covalently modified during catalysis offer a way to identify functional motions, as the modification can trigger catalytically important conformational changes.
The covalent modification of the active-site cysteine in isocyanide hydratase weakens a critical hydrogen bond required for reactivity. Hydrogen bond disruption triggers a cascade of conformational changes whose modulation by mutation is detrimental to enzyme turnover. Most enzymes that form catalytic intermediates will experience similar transient changes in active-site electrostatics, suggesting that modification-gated conformational dynamics is common in enzymes. Notably, all cysteine thiolates will experience a similar loss of negative charge upon covalent bond formation. Because cysteine residues can have multiple roles in a protein, including catalytic nucleophile, metal ligand, acylation target, redox target, and others, many different modification- mediated signals may be transduced through altered cysteine electrostatics to impact 99 protein dynamics and function, expanding the ways in which cysteine can couple protein biophysical properties to cellular needs. Integrating recent developments in serial and conventional RT crystallography with advanced computational methods531 can offer new, exciting opportunities to reveal enzyme mechanism and interrogate the consequences of cysteine modification on protein dynamics.
Modulation of protein dynamics is a powerful way to regulate protein function, as has been characterized in various systems. Cysteine modification-gated conformational changes in ICH alter the active site environment, promoting progress along the reaction coordinate. To our knowledge, ICH is the first example of a non-disulfide cysteine modification regulating functional protein conformational dynamics. Nevertheless, conceptually similar examples of gated conformational dynamical changes exist. Redox- gated changes in flavoprotein structure and dynamics may play a major role in electron transfer by these proteins558, and similar electron- or charge-coupled gating events occur in diverse systems 532,559,560. Photoactivatable tags that modulate sampling of active enzyme conformations have been used to create catalytically enhanced enzymes192. In the
- DJ-1 superfamily to which ICH belongs, C106 oxidation to C106-SO2 in DJ-1 results in little change in global protein conformation but stabilizes the protein by over 12 °C561.
This stabilization is thought to be due to a strong (2.47 Å) hydrogen bond between C106-
- SO2 and Glu18 that forms upon oxidation, reducing protein dynamics and stabilizing the protein. The many potential covalent modifications of cysteine make this residue of particular importance for understanding how cellular signaling states, metabolite pools, and stress conditions couple to functional protein dynamics through the modification of amino acids in proteins. Future work on ICH and other cysteine-containing proteins will 100 illuminate the diverse mechanisms by which cysteine-gated conformational changes can regulate protein function.
101
CHAPTER 5:
Purification, Kinetics and Structural Characterization of novel ICH homolog, RsICH from Ralstonia solanacearum GMI1000.
Note: The results described in this chapter have not been published.
Strained residues (in yellow spheres) cluster near functionally important regions in the RsICH dimer. The mobile helices are labeled “A” and “B” near the top of the dimer, and the active site C121 residues shown in sticks.
102
5.1 Abstract:
Isocyanide hydratase (ICH) catalyzes the conversion of isocyanide natural products to N-formamides by the addition of water. ICHs are widely distributed in pseudomonad bacteria, where they play a protective role against antimicrobial isocyanide natural products. We have previously characterized the ICH from Pseudomonas fluorescens (PfICH) using structural and biophysical approaches. PfICH exhibits collective motions that are activated by catalysis and permit water access to the catalytic thioimidate intermediate for hydrolysis. To determine if this dynamical aspect of ICH was a general feature, we recently determined an ultra-high resolution (0.74Å) X-ray crystal structure of wildtype ICH from Ralstonia solanacaerum GMI1000 (RsICH). Like
PfICH, RsICH exhibits backbone torsional strain near the active site cysteine residue that is important for helix motion in PfICH. In addition, RsICH also has an unusually large amount of side chain strain, including deviations from planarity of multiple residues near functionally important sites within the enzyme. Our working hypothesis is that fold- induced structural strain in the backbone and sidechains of these residues undergoes changes during catalysis to support correlated, functional motions. We expect that geometric stain in RsICH will relax upon intermediate formation and concurrent helix displacement and then increase as the catalytic cycle is completed. This chapter of the thesis will include ongoing and planned future work, involving X-ray crystallography, enzyme kinetics and site directed mutagenesis to determine if conformational strain in
RsICH is important for efficient catalysis. This model for the coupling between enzyme structure, dynamics, and function may provide general insight into how catalytic cysteine 103 covalent modification alters conformational ensembles and non-equilibrium motions in other cysteine-dependent enzymes.
5.2 Introduction:
Although enzyme catalysis is the central phenomenon of biochemistry, the relationship between their remarkable catalytic proficiency and structural and dynamical properties is not fully understood257,562,563. Because enzymes often catalyze complex reactions with one or more reaction intermediates, directly observing enzyme catalysis in real time and at atomic resolution has been a foundational goal of structural enzymology.
Technical advances in X-
ray crystallography now
permit the direct
monitoring of enzyme
turnover in crystalline
samples, including
identifying non-
equilibrium enzyme Figure 1: Clustal W pairwise sequence alignment. RsICH (WP_011001740.1) shares conserved catalytic residues with PfICH motions564–566 during (3NON) (36% identity). catalysis. These new technologies allow enzyme motions to be mapped while progressing along the reaction coordinate, addressing a long-standing challenge of visualizing and identifying functionally relevant enzyme dynamics. The primary focus of this thesis is cysteine- dependent ICH enzymes, which form covalent catalytic intermediates that modify the structural, electrostatic, and dynamic environment of the active site567,568. Catalysis- 104 activated enzyme motions provide a general model to explain how enzymes that use covalent catalysis are transiently modified to aid substrate turnover. Therefore, what is learned from ICH will be relevant to a large class of biomedically important enzymes.
The previous chapter (Chapter 4) discussed the formation of a catalytic cysteine (C101) covalent intermediate (thioimidate) in Pseudomonas ICH (PDB: 3NON). Formation of this cysteine covalent adduct alters active site electrostatics and induces dynamical changes that facilitate efficient substrate turnover and enzyme regeneration. To investigate how common cysteine-modification gated enzyme functional dynamics is, we identified a distant (36% identity) ICH homolog from Ralstonia solanacaerum GMI100
(NCBI Reference Sequence: WP_011001740.1) (RsICH) (Fig. 1).
5.3 Materials and Methods:
Cloning and Mutagenesis of Ralstonia solanacearum GMI1000 ICH (RsICH): The
RsICH gene was synthesized with E. coli codon optimization from Invitrogen (Thermo
Fisher Scientific) and inserted using Infusion cloning (Takara Bio USA, Inc) between the
NdeI and XhoI restriction sites of the bacterial expression vector pET15b (Novagen,
Darmstadt, Germany) so that the expressed protein carries an N-terminal, thrombin- cleavable hexa-histidine tag. The final recombinant RsICH protein is 224-amino acid long with a calculated molecular mass of 24,158 Da. All point mutations (E122Q,
E122D, E122T, C121S) were generated by site-directed mutagenesis using Q5 Site
Directed Mutagenesis (New England Labs) and High-Fidelity Phusion DNA polymerase
(Thermo Fisher Scientific) and appropriate mutagenic primers from Eurofins (Oligo synthesis). All ICH constructs were verified by DNA sequencing (Eurofins, Genewiz). 105
Protein Expression and Purification: E. coli BL21DE3 cells bearing the RsICH- pET15b construct were grown in LB medium supplemented with 100 μg/ml ampicillin, at
37 °C with shaking at 250 rpm until the A600 reached 0.5–0.7. Overexpression of ICH was induced by the addition of 0.66 mM isopropyl β-D-thiogalactopyranoside (IPTG), followed by 4 h of incubation at 37 °C. Cells were harvested by centrifugation, frozen in liquid nitrogen, and stored at −80 °C until needed. Frozen cell pellet was thawed on ice and resuspended in lysis buffer (50 mM HEPES, pH 7.5, 300 mM KCl, 10 mM imidazole). The suspended cells were lysed by the addition of hen egg white lysozyme (final concentration of 40 mg/ml) at 4 °C for 30 min, followed by sonication (5 second pulse on, 15 seconds pulse off settings) and centrifugation to remove cell debris.
The cleared supernatant (lysate) was applied to a His-select Ni2+ metal affinity column
(Sigma-Aldrich) that had been equilibrated in lysis buffer and bound for 1 hour at 4°C.
The recombinant hexa-histidine tagged RsICH was purified using Ni2+ metal affinity chromatography using 250 mM imidazole to elute bound protein. The eluted protein was dialyzed overnight at 4 °C against storage buffer (25 mM HEPES, pH 7.5, 100 mM KCl).
The following day, the hexa-histidine tag used for protein purification was removed by thrombin cleavage (2 units of thrombin/mg of ICH) at room temperature for 2-3 hrs, followed by passage over Ni2+ metal affinity resin to remove any uncleaved protein that retained the histidine tag. Thrombin was removed by passage over benzamidine-
Sepharose resin (GE Healthcare). The eluted protein was then dialyzed overnight at 4 °C against 25 mM HEPES, pH 7.5, 2mM DTT to remove any salts. The following day the protein was passed through UnoQ ion exchange chromatography (Bio-Rad). The protein fractions were pooled together, and protein concentration was determined using a 106
−1 −1 calculated extinction coefficient of ϵ280 = 31,065 M cm based on the amino acid composition of R. solanacearum ICH. The final protein shows up as a ∼24 kDa single band on overloaded Coomassie Blue-stained 12 % SDS-PAGE. Purified RsICH was concentrated to 16-27 mg/ml using a centrifugal concentrator (Amicon, Billerica, MA), supplemented with 2 mM DTT, frozen in liquid nitrogen, and stored at −80 °C. The
E122Q, E122D and E122T mutants were purified following the same protocol.
Crystallization, preliminary X-ray data collection, and processing: All crystals of
RsICH were grown at room temperature using the hanging drop vapor diffusion method.
For wild-type, E122Q, E122D, E122T RsICH, drops containing 2 μl of protein at 16-27 mg/ml and 2 μl of reservoir solution (16% PEG 3350, 0.1M MES, pH 6.5) were equilibrated against 500 μl of reservoir solution supplemented with 8% Dioxane or 8%
Ethanol. The choice of the additive is significant because 8% dioxane supports plate-like crystals, while 8% ethanol supports rod like crystals in both wildtype and the mutants.
Both plate type and rod type crystals were determined to be space group P 21 21 21. A third form of RsICH crystals were grown with 5-15% PEG 400 as additive; they were tetragonal in space group P 43 12 1.
All the crystals were cryo-protected by passage through six solutions of incrementally increasing amounts of ethylene glycol to a final concentration of 30% (v/v) in the reservoir solution. The cryoprotected crystal was cooled by immersion into liquid nitrogen. Preliminary wildtype and E122 D,Q and T RsICH crystal X-ray diffraction data at 1.5 Å were collected at the University of Nebraska Macromolecular Structural Core
Facility from single crystals maintained at 110 K using MicroMAX-007 rotating copper anode X-ray generator (Rigaku/MSC, The Woodlands, TX) operating at 40kV and 107
20mA, Osmic Blue confocal optics, and a Raxis IV++ image plate detector. All data were indexed, scaled, merged and refined using iMosflm, Aimless, Pointless, and
Refmac5 from the CCP4i suite, with final data statistics in Table 5.
RsICH microcrystal growth and fixed-target serial crystallography data collection and refinement: wt RsICH microcrystals were grown by batch seeding. Initial seeds were obtained by pulverizing macroscopic rod type crystals via vortexing with 0.5 mm stainless steel balls for 5-10 minutes. A 1:1500 dilution of this seed stock was added to
26% PEG 3350, 50 mM MES (pH 6.5), 16% Ethanol and 22% Ethylene glycol and then an equal volume of 38 mg/ml wtRsICH was added and rapidly mixed. The dilution of the seed stock dictated the final size of the microcrystals and was optimized to produce crystals measuring ~20x7x7 μm. The mixture was incubated at room temperature with shaking for ~20 minutes until microcrystal growth stopped. A 1.2 Å resolution serial crystallography dataset was collected at 100 K from microcrystals on a MiteGen mesh support using a fixed target strategy at SSRL beamline 12-2. This dataset comprised
~1400 indexed frames with 99.43 (95.62) % completeness, 23.36 (5.90) multiplicity, and a CC1/2 of 0.93. (0.17) (values for highest resolution shell in parentheses). This is a remarkably high-quality dataset given the limited number of images and the fact that it was collected at a synchrotron rather than an XFEL. The future goal is to collect a higher resolution MISC dataset at an XFEL due to the higher X-ray flux per exposure.
RsICH enzyme kinetics. The substrate used for all the kinetics experiments was para nitro-phenyl isocyanide (p-NPIC), synthesized by the Berkowitz group (University of
Nebraska Lincoln, Department of Chemistry). Steady state RsICH rate measurements were initiated by the addition of ICH (final concentration of 0.1 µM) to freshly prepared 108 p-NPIC solutions ranging from 0-1000 µM in reaction buffer (100 mM KPO4 pH=7.0, 50 mM KCl, and 20% DMSO, pH 7.6). p-NPIC was diluted from a 0.5 M stock solution in dimethyl sulfoxide (DMSO), which was stored at -80 °C and protected from light. A 200
µl reaction was maintained at 25 °C in a Peltier-thermostatted cuvette holder. The formation of the product, para-nitrophenyl formamide (p-NPF) was monitored at its absorption maximum of 320 nm for two minutes using a UV-Vis Cary 50
Spectrophotometer (Varian, Palo Alto, CA). A linear increase in A320 was verified and used to calculate initial velocities. The extinction coefficient at 320 nm for p-NPF was determined by using ICH to convert known concentrations of p-NPIC to p-NPF, followed by measuring the absorbance at 320 nm. The slope of the resulting standard curve was
4 -1 -1 defined as the extinction coefficient of p-NPF at 320 nm; ε320=1.33x10 M cm . This p-
NPF ε320 value was used to convert the measured rates from A320/sec to [p-NPF]/sec. All data were measured in triplicate or greater and mean values and standard deviations were plotted and fitted using the Michaelis-Menten model as implemented in Prism (GraphPad
Software, San Diego, CA).
Pre-steady state ICH kinetics were measured at 25 ˚C using Hi-Tech KinetAsyst stopped flow device (TgK Scientific, Bradford-on-Avon, United Kingdom). Data were collected for each sample for two seconds (instrument deadtime is 20 ms) in triplicates and product evolution was monitored at 320 nm using a photodiode array detector.
To ensure that the fixed target serial crystallography at SSRL beamline 12-2 was successful, as a pre-requisite, tests of catalysis were performed on RsICH microcrystals in the stabilization solution (26% PEG 3350, 50 mM MES (pH 6.5), 16% Ethanol and
22% Ethylene glycol). The crystals were pelleted at 7.5 rpm for 1 minute and then 109 resuspended in stabilization solution to remove any free protein. Rate measurements were performed by the addition of the wt RsICH microcrystals (final concentration of 5-50
µM) to freshly prepared p-NPIC solution at 500 µM in stabilization buffer (26% PEG
3350, 50 mM MES (pH 6.5), 16% Ethanol and 22% Ethylene glycol).Total reaction volume was maintained at 200 µl at 25 °C in a Peltier-thermostatted cuvette holder. The formation of the product, para-nitrophenyl formamide (p-NPF) was monitored at its absorption maximum of 320 nm for five minutes at 20 second intervals using a UV-Vis
Cary 50 Spectrophotometer (Varian, Palo Alto, CA).
Isocyanide (p-NPIC) toxicity assays. E coli BL21DE3 cells bearing the wtRsICH containing pET15b recombinant plasmids were grown until OD600=0.5-0.6 at 37˚C and
250rpm shaking, upon which they were induced with 0.5mM IPTG. 1 ml of induced cells were mixed rapidly with 5mls of cooled autoclaved soft agar (0.8%) supplemented with
100mg/ml Ampicillin and poured over 1.5% LB agar plates and incubated at room temperature (22-25˚C) for 1 hour. Once the soft agar had solidified, stress chemicals were prepared fresh (500mM, 100mM and 10mM p-NPIC in 100%DMSO). Each LBA plate was divided into four quadrants and sterile autoclaved filter discs (1 cm diameter) were placed in the four sections. 10µl stress chemical was added to each disk and the plates were incubated at 37˚C overnight. Diameters of the developed zone of clearances were manually measured in triplicates after 24 hours and plotted as a function of substrate concentration using Prism (GraphPad Software, San Diego, CA). The same protocol was followed for the mutants E122D, Q and T RsICH.
110
5.4 Results:
5.4.1. RsICH is a faster and more potent ICH compared to PfICH.
Like its homolog PfICH, Ralstonia ICH breaks down the isocyanide substrate p-
NPIC into the corresponding p-N formamide, which can be tracked spectrally at 320nm.
However, RsICH is much faster, with a turnover number of ~ 75 s-1compared to 0.2 s-1 in
PfICH (Fig. 9.A) (Tables 1,4). Pre-steady state kinetics of RsICH indicates an absence of a burst phase that is present with PfICH (Fig. 9 D-C). This indicates that the process of thioimidate hydrolysis and
Figure 2: (A). Impaired steady state Michelis Menten kinetics of (clockwise from left) wt, E122D, Q, T RsICH. (B). E. coli BL21DE3 p-NPIC toxicity assay reveals in vivo effects of impaired catalysis in E122 mutants. (C). Disk diffusion assay showing zones of inhibited E. coli growth near disks impregnated with the indicated concentrations (mM) of p-NPIC. (D). PfICH shows burst phase in stopped flow mixing experiment. (E). RfICH exhibits no burst phase, indicating faster rates of thioimidate formation/hydrolysis
111 product formation in RsICH must occur faster than in PfICH and the rates of formation and hydrolysis are more closely matched or that the rate-limiting step in RsICH is now thioimidate formation. It is possible that RsICH also has a burst phase that is within the
~20 ms dead time of the instrument, although this seems unlikely. To test this hypothesis, we could perform the stopped flow experiment in a reaction buffer in which RsICH is slower to lengthen the pre steady-state rate. One potential way to do this is by optimizing the reaction buffer composition. We have measured catalytic turnover in microcrystals of
RsICH in the crystallization buffer (16% PEG 3350, 0.5M MES/Citrate pH 6.5, 8% ethanol) (Fig. 6.B) (More details in section 5.4.4). The rate of turnover can be controlled by changing the relative concentrations of the MES or citrate buffers (both at pH 6.5) in the crystal (Fig. 6.D). MES decreases in crystallo reaction rates because it is loosely bound at a site that partially occludes a solvent channel to the active site in the 0.74 Å resolution crystal structure (Fig. 6.C). Although these experiments have not been conducted with the enzyme in solution, it will be worth investigating if tuning the MES: citrate ratio in the reaction buffer sufficiently slows down RsICH in solution catalysis to observe the intermediate formation/hydrolysis step. Another possible method would be to employ single-turnover kinetics, which would allow further isolation of the first catalytic turnover of the enzyme
5.4.2. E122 D, Q and T mutations in the RsICH active site significantly impairs catalysis. E122 residue, immediately following the reactive C121, was mutated to a
Threonine (E122T) to simulate the active site environment of the Pseudomonas ICH, to investigate the contribution of E122 to the faster rates of catalysis in RsICH. E122D mutant was generated to investigate the potential steric effects of E122 within the active 112 site, and finally, the E122Q mutant was made to disrupt the hydrogen bonding environment within the active site. The E122 D, Q, T mutants exhibit impaired steady state kinetics (Fig.2.A) (Table 4) with turnover numbers ~ 0.2 s-1, 0.4 s-1 and 0.3 s-
1respectively. p-NPIC toxicity assays performed on E. coli BL21DE3 cells bearing the wildtype RsICH construct show that these cells tolerate isocyanide much better than E. coli cells expressing PfICH (Fig. 2. B-C). The E122D, Q and T RsICH mutations, however, diminish isocyanide tolerance levels to
Figure 3: (A-B). 0.74 Å dataset of RsICH. 2Fo-Fc electron density (blue 1.0σ) and Fo-Fc difference density (green +3σ, red -3σ) show deprotonated bond lengths of E122 (1.24-1.25 Å) and 3.48 Å hydrogen bond with C121 thiolate. (C) conserved C147-C220 disulfide bond in RsICH (2.08 Å). comparable with wt and G150A and G150T PfICH (Fig. 5, Chapter 4).
Although the E122 mutation seems to significantly impair catalysis, it is not yet obvious why. Neither is it clear why RsICH is a faster ICH than PfICH. In the 113
Pseudomonas ICH, the reactive C101 (PfICH numbering) is immediately followed by a
T102, while in Ralstonia ICH it is a Glu122 (RsICH numbering). Moreover, ultra-high resolution (0.74-0.9 Å) resolution dataset(s) of RsICH crystals reveal that E122 is deprotonated (hence negatively charged) with a Cδ-Oε1/2 bond lengths of ~1.25 Å
(Fig.10. A) (Data statistics in Table 6). Surprisingly, this ionized carboxylate is 3.48 Å away from the reactive C121 (RsICH numbering) in wt and E122D, Q, T RsICH. We know that for the catalytic reaction to occur, C121 must be a nucleophile, thus deprotonated to the thiolate. It is unclear how these two negatively charged neighboring residues (E122, C121) would stably exist so near to one another in the active site440
(Fig.10. A-B). It is important to note here that a protein BLAST revealed several ICH homologs with a glutamic acid following the nucleophilic Cys; however it remains to be examined whether these glutamic acids are also deprotonated as seen in RsICH.
It is possible that the negatively charged electrostatic environment created by both
C121 and E122 interacts with the formal positive charge on the nitrogen atom of the isocyanide substrate and helps position the substrate for attack in the active site of the enzyme. In this case, RsICH has greater potential for electrostatically pre-organizing the
Michaelis complex than the slower PfICH, which has a threonine (T102) (Pseudomonas numbering) at this position. It is also possible that the strong negative charge created by these two residues in RsICH electrostatically strains the active site prior to catalysis, and that formation of the thioimidate neutralizes the negative charge of C121 relieves this electrostatic strain and stabilizes the Michaelis complex. The restoration of the thiolate upon hydrolysis of the thioimidate may restore electrostatic strain that facilitates product release. Another hypothesis is that C121 is initially protonated as a thiol and binding of 114 the substrate p-NPIC leads to its ionization; the deprotonated C121 nucleophile then attacks the electrophilic isocyanide carbon and initiates the reaction. Additional experimental work is required to experimentally test these hypotheses. It would be interesting to see if making a T102E mutation in PfICH (PfICH numbering) speeds up the reaction to similar rates as seen in RsICH.
Another unusual structural feature observed in RsICH is the existence of a conserved disulfide bond between two of its cysteines (C147 and C220) near the C- terminus of the enzyme (Fig.10. C), even though RsICH must folded in the reducing
E.coli cytosol during recombinant protein expression573,574 . The high degree of conservation of the cysteine residues in this bond, as well as its robustness to DTT
(1mM) to prevent oxidation during protein purification both suggest possible functional importance. Because disulfide bonds are typically formed and maintained within an oxidizing environment like the endoplasmic reticulum in eukaryotes574, one possibility is that the C147-C220 disulfide bond is formed in the more oxidizing E.coli periplasm 422.
However, proteins are targeted into the periplasm by specific signal sequences575,575–577 and it remains to be explored whether RsICH is targeted in a similar manner.
5.4.3. RsICH exhibits cysteine-gated active site helix dynamics similar to PfICH.
We have obtained a 1.5 Å resolution rotating anode crystal structure of RsICH at pH 5.5
(Table 5). This lower pH structure shows clear evidence of crystallographic disorder near the highly strained Ramachandran outlier I175 (corresponding to I152 in Pseudomonas
ICH), indicating 115
Figure 4: (A). Ralstonia ICH shows mobility near the active site at pH 5.5. 2Fo-Fc electron density (blue 1.0σ) and Fo-Fc difference density (green +3σ, red -3σ) show clear evidence of two conformations of the active site C121 (A, B) and mobility of the strained I175 in the mobile helix. (B) No evidence of helix mobility in Ralstonia ICH pH 6.5. 2Fo-Fc electron density (blue 1.0σ) and Fo-Fc difference density (green +3σ, red -3σ). helical mobility (Fig. 4). We observe this disorder at pH 5.5 and not in the 0.74 Å resolution structure at pH 6.5 likely because the active site cysteine is protonated at 5.5, neutralizing its charge and weakening the H-bond with I175. This is the same charge- gated H-bond mechanism that we propose for Pseudomonas ICH in chapter 4.
5.4.4. Identification of side chain geometric strain in the ultrahigh resolution crystal structure of RsICH.
Figure 5: (A) 2Fo-Fc electron density (blue 1.0σ; purple 4.5σ) at 0.74 Å resolution for W85 shows the exceptional quality of the data. The protrusions at in the 2Fo-Fc electron density are hydrogen atoms. (B) Side view of W85 shows obvious deviations from planarity. (C) Strained residues (in yellow spheres) cluster near functionally important egions in the Ralstonia ICH dimer; the mobile helices are labeled “A” and “B” near the top of the dimer, and the active site C121 residues shown in sticks. 116
We have recently collected a synchrotron X-ray diffraction dataset of Ralstonia ICH
(RsICH) at 100 K to 0.74 Å resolution (Table 6). This ultrahigh resolution dataset allows the well-ordered parts of the structure to be refined with no influence from geometric restraints, as the observation-to-parameter ratio for unrestrained refinement is ~13. Even when restraints are maintained, residues Y15, W37, W85, R162, Y163, and Y204 all exhibit pronounced planarity deviations. W85 in particularly is extremely distorted, with a clearly nonplanar indole ring (Fig. 5). In Ralstonia ICH (RsICH), most of the strained residues cluster near the active site or surround the helix that is mobile during catalysis in
Pseudomonas ICH (PfICH). It is possible that this conformational strain in RsICH facilitates important conformational changes in the enzyme during catalysis, and as such, we strongly suspect a connection between these strained residues and RsICH catalysis.
5.4.5. Future work will include tracking the origin and dynamics of conformational strain in RsICH during catalysis using MISC.
Future work will involve the use of computational methods to identify the origin of the strain in the mentioned residues, allowing us to determine the physical origin of these effects, including the energetics of distortion and the major structural determinants of these deviations. Building from the computational results, we will mutate those residues identified as distorting the strained sidechains in order to reduce or eliminate strain. We will determine crystal structures of these designed mutants to validate the effects on geometric strain and then compare computationally predicted and observed sidechain geometries. Using these strain-modulating mutants, we will then perform steady and pre steady-state kinetic analysis using p-NPIC as substrate. By combining 117 computational, structural, and kinetics approaches, we will be able to determine the contribution of sidechain strain to Ralstonia ICH (RsICH) catalytic proficiency.
We will also perform accurate measurement of the geometric strain in the non- planar residues in RsICH. For this, we will need atomic resolution (≤1.2 Å) data for
Ralstonia ICH, which will be the highest resolution MISC dataset ever collected.
Preliminarily, we have established the feasibility of this experiment by reliably growing microcrystals (~20x7x7 μm) (Fig. 6.A) and collecting a 1.2 Å resolution serial crystallography dataset from a fixed target mesh at SSRL beamline 12-2 (Methods).
These high-resolution time-resolved data will be used to investigate the changes in geometric strain that we hypothesize may facilitate motions that are important for turnover. This could potentially be the first experiment in which conformational strain will be directly observed in an enzyme during catalysis. To establish feasibility of the planned MISC component of the experiment, we measured catalytic turnover in microcrystals of Ralstonia ICH in the crystallization buffer (16% PEG 3350, 0.5M
MES/Citrate pH 6.5, 8% ethanol) (Fig. 6.B). The rate of turnover can be tuned by changing the relative concentrations of the MES or citrate buffers (both at pH 6.5) in the crystal (Fig. 6.D). MES decreases in crystallo reaction rates because it is loosely bound at a site that partially occludes a solvent channel to the active site in the 0.74 Å resolution crystal structure (Fig. 6.C). 118
Figure 6: (A) RsICH microcrystals. (B) Ralstonia ICH microcrystals are catalytically active. Spectra were collected every 20 s, from purple to red. The product absorbs at 320 nm. (C) MES bound in the active site of RsICH. 2Fo-Fc electron density (blue 1.0σ) and Fo-Fc difference density (green +3σ, red -3σ) (D) Rates of microcrystal catalysis can be altered by manipulating the Citrate: MES (pH 6.5) ratio in crystallization buffer.
We expect that Ralstonia ICH will exhibit catalysis-gated motions that are coupled to intermediate hydrolysis, with geometric strain relaxing during intermediate formation/helix displacement and then increasing again as the catalytic cycle is completed. Importantly, the atomic resolution of the data will make this a first-of-kind
MISC experiment, and the resulting data will allow the development of improved tools for all steps in the serial crystallography data processing and model refinement workflow that will benefit the structural biology community.
119
5.5 Discussions:
Preliminary kinetics and in vivo toxicity assays show that Ralstonia ICH (RsICH) is a faster ICH enzyme compared to its Pseudomonas (PfICH) homolog. Moreover,
RsICH diffracts to ultra-high resolution (0.74 Å) and diffracts to atomic resolution (≤1.2
Å) in a serial crystallography experiment. There are only about a dozen ultra-high resolution datasets (≤0.07 Å) in existence 582. Interestingly, the 0.74Å RsICH dataset shows fold-induced backbone and sidechain strain. Particularly, amino acid residues
Y15, W37, W85, R162, Y163, and Y204 all exhibit pronounced planarity deviations, which is highly unexpected. Other ultrahigh resolution datasets also exhibit aromatic planarity deviations, and these strained residues are typically in functionally important parts of those structures569–572. In RsICH, most of the strained residues cluster near the active site or surround the helix that is mobile during catalysis in PfICH, strongly suggesting a connection to ICH function, which remains to be explored.
Because of the exceptional resolution of the preliminary synchrotron X-ray dataset (0.74 Å), these deviations from planarity are well-determined by the data and thus residue geometry is confidently known. Similar geometric strain has been observed in other proteins that diffract to ultrahigh resolution583,584, but its origin is unknown. Future work will include determination of the origin of this strain and investigate how catalytic modification of the active site couples to strained residues throughout the protein to activate functional motions. The proposed work will elucidate general physical mechanisms by which cysteine modification alters protein structure and function, with potentially wide applicability to other biomedical important proteins, including the development of small molecules that covalently target key cysteine residues. 120
Works cited
1. Lieke, W. Ueber das Cyanallyl. Justus Liebigs Annalen der Chemie 112, 316–321.
2. Edenborough, M. S. & Herbert, R. B. Naturally occurring isocyanides. Nat Prod
Rep 5, 229–245 (1988).
3. Gautier, A. Ueber die Carbylamine; Justus Liebigs Annalen der Chemie 149, 29–35
(1869).
4. Keating, T. A. & Armstrong, R. W. Postcondensation Modifications of Ugi Four-
Component Condensation Products: 1-Isocyanocyclohexene as a Convertible
Isocyanide. Mechanism of Conversion, Synthesis of Diverse Structures, and
Demonstration of Resin Capture. J. Am. Chem. Soc. 118, 2574–2583 (1996).
5. Kemp, D. S. & Woodward, R. B. The N-ethylbenzisoxazolium cation—I:
Preparation and reactions with nucleophilic species. Tetrahedron 21, 3019–3035
(1965).
6. Kreutzkamp, N. & Lämmerhirt, K. Phosphinylmethyl Isocyanides. Angewandte
Chemie International Edition in English 7, 372–373 (1968).
7. Dömling, A. Recent Developments in Isocyanide Based Multicomponent Reactions
in Applied Chemistry. Chem. Rev. 106, 17–89 (2006).
8. Belluco, U., Crociani, B., Michelin, R. & Uguagliati, P. Electrophilic and
nucleophilic attacks on alkyl, nitrile, and isonitrile complexes of Pd(II) and Pt(II).
Pure and Applied Chemistry 55, 47–54 (1983).
9. Singleton, E. & Oosthuizen, H. E. Metal Isocyanide Complexes. in Advances in
Organometallic Chemistry (eds. Stone, F. G. A. & West, R.) vol. 22 209–310
(Academic Press, 1983). 121
10. Witczak, Z. J. Review Article Monosaccharide Isocyanides, Synthesis, Chemistry
and Application. Journal of Carbohydrate Chemistry 3, 359–380 (1984).
11. Ugi, I. & Wischhöfer, E. Isonitrile, XI. Synthese einfacher Penicillansäure-Derivate.
Chemische Berichte 95, 136–140 (1962).
12. Bredereck, H., Föhlisch, B. & Walz, K. Reaktionen der N-Isocyan-dialkylamine.
Justus Liebigs Annalen der Chemie 688, 93–98 (1965).
13. Nef, J. U. Ueber das zweiwerthige Kohlenstoffatom. Justus Liebigs Annalen der
Chemie 270, 267–335.
14. Lindemann, H. & Wiegrebe, L. Über die Konstitution der Verbindungen mit
„zweiwertigem” Kohlenstoff. Berichte der deutschen chemischen Gesellschaft (A
and B Series) 63, 1650–1657.
15. Dadieu, A. & Kohlrausch, K. W. F. Raman-Effekt und Chemie. Berichte der
deutschen chemischen Gesellschaft (A and B Series) 63, 251–282.
16. Harris, N. C. et al. Isonitrile Formation by a Non-Heme Iron(II)-Dependent
Oxidase/Decarboxylase. Angew. Chem. Int. Ed. Engl. (2018)
doi:10.1002/anie.201804307.
17. Rothe, W. [The new antibiotic xanthocillin.]. Dtsch Med Wochenschr 79, 1080–
1081 (1954).
18. Hagedorn, I. & Tonjes, H. [Structure of xanthocillin, a new antibiotic]. Pharmazie
11, 409–410 (1956).
19. Scheuer, P. J. Isocyanides and cyanides as natural products. Acc. Chem. Res. 25,
433–439 (1992). 122
20. Cafieri, F., Fattorusso, E., Magno, S., Santacroce, C. & Sica, D. Isolation and
structure of axisonitrile-1 and axisothiocyanate-1 two unusual sesquiterpenoids from
the marine spongeAxinella cannabina. Tetrahedron 29, 4259–4262 (1973).
21. Puar, M. S., Munayyer, H., Hegde, V., Lee, B. K. & Waitz, J. A. THE
BIOSYNTHESIS OF HAZIMICINS: POSSIBLE ORIGIN OF ISONITRILE
CARBON. J. Antibiot. 38, 530–532 (1985).
22. M. Cable, K., B. Herbert, R., R. Knaggs, A. & Mann, J. The biosynthesis of
xanthocillin-X monomethyl ether in dichotomomyces cejpii . Experiments on the
origin of the isocyanide carbon atoms. Journal of the Chemical Society, Perkin
Transactions 1 0, 595–599 (1991).
23. Cable, K. M., Herbert, R. B. & Mann, J. On the biosynthesis of the fungal
isocyanide, xanthocillin monomethyl ether. Tetrahedron Letters 28, 3159–3162
(1987).
24. Achenbach, H. & König, F. Zur Biogenese des Xanthocillins, III Die Frage der
biogenetischen Gleichwertigkeit der beiden Xanthocillin-Hälften. Chemische
Berichte 105, 784–793 (1972).
25. E. Baldwin, J. et al. Biosynthesis of a cyclopentyl dienyl isonitrile acid in cultures
of the fungus Trichoderma hamatum (Bon.) bain. aggr. Journal of the Chemical
Society, Chemical Communications 0, 1227–1229 (1981).
26. Parry, R. J. & Buu, H. P. Biosynthesis of an isonitrile acid from
trichodermahamatum. Tetrahedron Letters 23, 1435–1438 (1982).
27. B. Herbert, R. & Mann, J. Biosynthesis of tuberin from tyrosine and glycine.
Journal of the Chemical Society, Chemical Communications 0, 1008–1010 (1983). 123
28. Bornemann, Volker., Patterson, G. M. L. & Moore, R. E. Isonitrile biosynthesis in
the cyanophyte Hapalosiphon fontinalis. J. Am. Chem. Soc. 110, 2339–2340 (1988).
29. Chang, C. W. J. Naturally Occurring Isocyano/Isothiocyanato and Related
Compounds. in Fortschritte der Chemie organischer Naturstoffe / Progress in the
Chemistry of Organic Natural Products (eds. Chang, C. W. J. et al.) 1–186
(Springer Vienna, 2000). doi:10.1007/978-3-7091-6331-3_1.
30. Pfeifer, S., Bär, H. & Zarnack, J. [Metabolites of the xanthocillin(Brevicid)-
producing mutant of Penicillium notatum Westl]. Pharmazie 27, 536–542 (1972).
31. Lakshminarasimhan, M., Madzelan, P., Nan, R., Milkovic, N. M. & Wilson, M. A.
Evolution of new enzymatic function by structural modulation of cysteine reactivity
in Pseudomonas fluorescens isocyanide hydratase. J. Biol. Chem. jbc.M110.147934
(2010) doi:10.1074/jbc.M110.147934.
32. Goda, M., Hashimoto, Y., Shimizu, S. & Kobayashi, M. Discovery of a Novel
Enzyme, Isonitrile Hydratase, Involved in Nitrogen-Carbon Triple Bond Cleavage.
J. Biol. Chem. 276, 23480–23485 (2001).
33. Fukatsu, H., Hashimoto, Y., Goda, M., Higashibata, H. & Kobayashi, M. Amine-
synthesizing enzyme N-substituted formamide deformylase: Screening, purification,
characterization, and gene cloning. PNAS 101, 13726–13731 (2004).
34. Brady, S. F. & Clardy, J. Cloning and Heterologous Expression of Isocyanide
Biosynthetic Genes from Environmental DNA. Angewandte Chemie 117, 7225–
7227 (2005). 124
35. Brady, S. F. & Clardy, J. Systematic Investigation of the Escherichia coli
Metabolome for the Biosynthetic Origin of an Isocyanide Carbon Atom.
Angewandte Chemie 117, 7207–7210 (2005).
36. Chang, W. et al. In Vitro Stepwise Reconstitution of Amino Acid Derived Vinyl
Isocyanide Biosynthesis: Detection of an Elusive Intermediate. Org. Lett. 19, 1208–
1211 (2017).
37. Brady, S. F., Bauer, J. D., Clarke-Pearson, M. F. & Daniels, R. Natural Products
from isnA-Containing Biosynthetic Gene Clusters Recovered from the Genomes of
Cultured and Uncultured Bacteria. J. Am. Chem. Soc. 129, 12102–12103 (2007).
38. Stratmann, K. et al. Welwitindolinones, Unusual Alkaloids from the Blue-Green
Algae Hapalosiphon welwitschii and Westiella intricata. Relationship to
Fischerindoles and Hapalinodoles. Journal of the American Chemical Society 116,
9935–9942 (1994).
39. Hillwig, M. L., Zhu, Q. & Liu, X. Biosynthesis of Ambiguine Indole Alkaloids in
Cyanobacterium Fischerella ambigua. ACS Chem. Biol. 9, 372–377 (2014).
40. Mo, S., Krunic, A., Chlipala, G. & Orjala, J. Antimicrobial Ambiguine Isonitriles
from the Cyanobacterium Fischerella ambigua. J. Nat. Prod. 72, 894–899 (2009).
41. Raveh, A. & Carmeli, S. Antimicrobial Ambiguines from the Cyanobacterium
Fischerella sp. Collected in Israel. J. Nat. Prod. 70, 196–201 (2007).
42. Hillwig, M. L. et al. Identification and Characterization of a Welwitindolinone
Alkaloid Biosynthetic Gene Cluster in the Stigonematalean Cyanobacterium
Hapalosiphon welwitschii. ChemBioChem 15, 665–669 (2014). 125
43. Stintzi, A. et al. The pvc Gene Cluster of Pseudomonas aeruginosa: Role in
Synthesis of the Pyoverdine Chromophore and Regulation by PtxR and PvdS. J
Bacteriol 181, 4118–4124 (1999).
44. Drake, E. J. & Gulick, A. M. Three-dimensional Structures of Pseudomonas
aeruginosa PvcA and PvcB, Two Proteins Involved in the Synthesis of 2-Isocyano-
6,7-dihydroxycoumarin. Journal of Molecular Biology 384, 193–205 (2008).
45. Clarke-Pearson, M. F. & Brady, S. F. Paerucumarin, a New Metabolite Produced by
the pvc Gene Cluster from Pseudomonas aeruginosa. Journal of Bacteriology 190,
6927–6930 (2008).
46. Gross, H. & Loper, J. E. Genomics of secondary metabolite production by
Pseudomonas spp. Nat. Prod. Rep. 26, 1408–1446 (2009).
47. Hamood, A. N., Colmer, J. A., Ochsner, U. A. & Vasil, M. L. Isolation and
characterization of a Pseudomonas aeruginosa gene, ptxR, which positively
regulates exotoxin A production. Molecular Microbiology 21, 97–110 (1996).
48. Carty, N. L. et al. PtxR modulates the expression of QS-controlled virulence factors
in the Pseudomonas aeruginosa strain PAO1. Molecular Microbiology 61, 782–794
(2006).
49. Tørring, T., Shames, S. R., Cho, W., Roy, C. R. & Crawford, J. M. Acyl Histidines:
New N-Acyl Amides from Legionella pneumophila. ChemBioChem 18, 638–646
(2017).
50. Faucher, S. P., Mueller, C. A. & Shuman, H. A. Legionella Pneumophila
Transcriptome during Intracellular Multiplication in Human Macrophages. Front
Microbiol 2, 60 (2011). 126
51. Crawford, J. M., Portmann, C., Zhang, X., Roeffaers, M. B. J. & Clardy, J. Small
molecule perimeter defense in entomopathogenic bacteria. PNAS 109, 10821–10826
(2012).
52. Takahashi, S. et al. Byelyankacin: A Novel Melanogenesis Inhibitor Produced by
Enterobacter sp. B20. The Journal of Antibiotics 60, 717–720 (2007).
53. Harris, N. C. et al. Biosynthesis of isonitrile lipopeptides by conserved
nonribosomal peptide synthetase gene clusters in Actinobacteria. PNAS 114, 7025–
7030 (2017).
54. Wang, L. et al. Diisonitrile Natural Product SF2768 Functions As a Chalkophore
That Mediates Copper Acquisition in Streptomyces thioluteus. ACS Chem. Biol. 12,
3067–3075 (2017).
55. Zhu, M. et al. Tandem Hydration of Diisonitriles Triggered by Isonitrile Hydratase
in Streptomyces thioluteus. Org. Lett. 20, 3562–3565 (2018).
56. Lim, F. Y. et al. Fungal Isocyanide Synthases and Xanthocillin Biosynthesis in
Aspergillus fumigatus. mBio 9, e00785-18 (2018).
57. Briza, P., Eckerstorfer, M. & Breitenbach, M. The sporulation-specific enzymes
encoded by the DIT1 and DIT2 genes catalyze a two-step reaction leading to a
soluble LL-dityrosine-containing precursor of the yeast spore wall. PNAS 91, 4524–
4528 (1994).
58. Briza, P., Kalchhauser, H., Pittenauer, E., Allmaier, G. & Breitenbach, M. N,N′
Bisformyl Dityrosine is an in vivo Precursor of the Yeast Ascospore Wall.
European Journal of Biochemistry 239, 124–131 (1996). 127
59. Qaisar, U. et al. The pvc Operon Regulates the Expression of the Pseudomonas
aeruginosa Fimbrial Chaperone/Usher Pathway (Cup) Genes. PLOS ONE 8, e62735
(2013).
60. Sivaneson, M., Mikkelsen, H., Ventre, I., Bordi, C. & Filloux, A. Two-component
regulatory systems in Pseudomonas aeruginosa: an intricate network mediating
fimbrial and efflux pump gene expression. Molecular Microbiology 79, 1353–1366
(2011).
61. Hindre, T., Bruggemann, H., Buchrieser, C. & Hechard, Y. Transcriptional profiling
of Legionella pneumophila biofilm cells and the influence of iron on biofilm
formation. Microbiology 154, 30–41 (2008).
62. Banin, E., Vasil, M. L. & Greenberg, E. P. Iron and Pseudomonas aeruginosa
biofilm formation. PNAS 102, 11076–11081 (2005).
63. Lin, M.-H., Shu, J.-C., Huang, H.-Y. & Cheng, Y.-C. Involvement of Iron in
Biofilm Formation by Staphylococcus aureus. PLoS One 7, (2012).
64. Johnson, M., Cockayne, A., Williams, P. H. & Morrissey, J. A. Iron-Responsive
Regulation of Biofilm Formation in Staphylococcus aureus Involves Fur-Dependent
and Fur-Independent Mechanisms. Journal of Bacteriology 187, 8211–8215 (2005).
65. Kang, D., Kirienko, D. R., Webster, P., Fisher, A. L. & Kirienko, N. V. Pyoverdine,
a siderophore from Pseudomonas aeruginosa, translocates into C. elegans, removes
iron, and activates a distinct host response. Virulence 9, 804–817 (2018).
66. Musk, D. J., Banko, D. A. & Hergenrother, P. J. Iron Salts Perturb Biofilm
Formation and Disrupt Existing Biofilms of Pseudomonas aeruginosa. Chemistry &
Biology 12, 789–796 (2005). 128
67. Oglesby-Sherrouse, A. G., Djapgne, L., Nguyen, A. T., Vasil, A. I. & Vasil, M. L.
The complex interplay of iron, biofilm formation, and mucoidy affecting
antimicrobial resistance of Pseudomonas aeruginosa. Pathog Dis 70, 307–320
(2014).
68. Patriquin, G. M. et al. Influence of Quorum Sensing and Iron on Twitching Motility
and Biofilm Formation in Pseudomonas aeruginosa. Journal of Bacteriology 190,
662–671 (2008).
69. Wilderman, P. J. et al. Identification of tandem duplicate regulatory small RNAs in
Pseudomonas aeruginosa involved in iron homeostasis. PNAS 101, 9792–9797
(2004).
70. Treichel, P. M. Transition Metal-lsocyanide Complexes. in Advances in
Organometallic Chemistry (eds. Stone, F. G. A. & West, R.) vol. 11 21–86
(Academic Press, 1973).
71. Mokhtarzadeh, C. C. et al. Synthesis and Protonation of an Encumbered Iron
Tetraisocyanide Dianion. Inorg. Chem. 54, 5579–5587 (2015).
72. Makino, R. et al. Oxygen Binding and Redox Properties of the Heme in Soluble
Guanylate Cyclase: IMPLICATIONS FOR THE MECHANISM OF LIGAND
DISCRIMINATION. Journal of Biological Chemistry 286, 15678–15687 (2011).
73. Mims, M. P., Porras, A. G., Olson, J. S., Noble, R. W. & Peterson, J. A. Ligand
binding to heme proteins. An evaluation of distal effects. J. Biol. Chem. 258,
14219–14232 (1983). 129
74. Rustad, T. R. et al. Mapping and manipulating the Mycobacterium tuberculosis
transcriptome using a transcription factor overexpression-derived regulatory
network. Genome Biology 15, 502 (2014).
75. Botella, H. et al. Mycobacterial P1-Type ATPases Mediate Resistance to Zinc
Poisoning in Human Macrophages. Cell Host & Microbe 10, 248–259 (2011).
76. Ueda, K., Tomaru, Y., Endoh, K. & Beppu, T. Stimulatory Effect of Copper on
Antibiotic Production and Morphological Differentiation in Streptomyces
tanashiensis. J. Antibiot. 50, 693–695 (1997).
77. R. Worrall, J. A. & Vijgenboom, E. Copper mining in Streptomyces : enzymes,
natural products and development. Natural Product Reports 27, 742–756 (2010).
78. Noguchi, A., Kitamura, T., Onaka, H., Horinouchi, S. & Ohnishi, Y. A copper-
containing oxidase catalyzes C-nitrosation in nitrosobenzamide biosynthesis. Nature
Chemical Biology 6, 641–643 (2010).
79. Wiemann, P. et al. Aspergillus fumigatus Copper Export Machinery and Reactive
Oxygen Intermediate Defense Counter Host Copper-Mediated Oxidative
Antimicrobial Offense. Cell Reports 19, 1008–1021 (2017).
80. Cross, A. R., Parkinson, J. F. & Jones, O. T. G. The superoxide-generating oxidase
of leucocytes. NADPH-dependent reduction of flavin and cytochrome b in
solubilized preparations. Biochemical Journal 223, 337–344 (1984).
81. Dershwitz, M. & Novak, R. F. Generation of superoxide via the interaction of
nitrofurantoin with oxyhemoglobin. J. Biol. Chem. 257, 75–79 (1982).
82. Ding, C., Festa, R. A., Sun, T.-S. & Wang, Z.-Y. Iron and copper as virulence
modulators in human fungal pathogens. Molecular Microbiology 93, 10–23 (2014). 130
83. Djoko, K. Y., Ong, C. Y., Walker, M. J. & McEwan, A. G. Copper and zinc toxicity
and its role in innate immune defense against bacterial pathogens. J. Biol. Chem.
jbc.R115.647099 (2015) doi:10.1074/jbc.R115.647099.
84. Percival, S. S. Neutropenia Caused by Copper Deficiency: Possible Mechanisms of
Action. Nutr Rev 53, 59–66 (1995).
85. Babu, U. & Failla, M. L. Respiratory Burst and Candidacidal Activity of Peritoneal
Macrophages Are Impaired in Copper-Deficient Rats. J Nutr 120, 1692–1699
(1990).
86. Muñoz, C., Rios, E., Olivos, J., Brunser, O. & Olivares, M. Iron, copper and
immunocompetence. British Journal of Nutrition 98, S24–S28 (2007).
87. Zaidi, K. U., Ali, S. A. & Ali, A. S. Effect of Purified Mushroom Tyrosinase on
Melanin Content and Melanogenic Protein Expression. Biotechnology Research
International https://www.hindawi.com/journals/btri/2016/9706214/ (2016)
doi:10.1155/2016/9706214.
88. Eleftherianos, I. et al. An antibiotic produced by an insect-pathogenic bacterium
suppresses host defenses through phenoloxidase inhibition. Proceedings of the
National Academy of Sciences of the United States of America 104, 2419–2424
(2007).
89. Kim, J.-P. et al. Melanocins A, B and C, New Melanin Synthesis Inhibitors
Produced by Eupenicillium shearii. J. Antibiot. 56, 993–999 (2003).
90. Lee, C. H. et al. MR566A and MR566B, New Melanin Synthesis Inhibitors
Produced by Trichoderma harzianum. J. Antibiot. 50, 474–478 (1997). 131
91. Tsuchiya, T. et al. Purification and Determination of the Chemical Structure of the
Tyrosinase Inhibitor Produced by Trichoderma viride Strain H1-7 from a Marine
Environment. Biological & Pharmaceutical Bulletin 31, 1618–1620 (2008).
92. Kozlovsky, A. G. et al. Penicillium expansum, a Resident Fungal Strain of the
Orbital Complex Mir, Producing Xanthocyllin X and Questiomycin A. Applied
Biochemistry and Microbiology 40, 291–295 (2004).
93. Kozlovskii, A. G. et al. The Biosynthesis of Low-Molecular-Weight Nitrogen-
Containing Secondary Metabolites—Alkaloids—by the Resident Strains of
Penicillium chrysogenum and Penicillium expansum Isolated on Board the Mir
Space Station. Microbiology 71, 666–669 (2002).
94. Yamamoto, Y. & Arai, K. Chapter 4 Alkaloidal Substances from Aspergillus
Species. in The Alkaloids: Chemistry and Pharmacology vol. 29 185–263 (Elsevier,
1986).
95. Takatsuki, A., Suzuki, S., Ando, K., Tamura, G. & Arima, K. Studies on Antiviral
and Antitumor Antibiotics. Agricultural and Biological Chemistry 33, 1119–1969
(1969).
96. Büchi, G., Luk, K. C., Kobbe, B. & Townsend, J. M. Four new mycotoxins of
Aspergillus clavatus related to tryptoquivaline. J. Org. Chem. 42, 244–246 (1977).
97. Kitahara, N. & Endo, A. Xanthocillin X monomethyl ether, a potent inhibitor of
prostaglandin biosynthesis. J. Antibiot. 34, 1556–1561 (1981).
98. Takahashi, C., Sekita, S., Yoshihira, K. & Natori, S. The structures of toxic
metabolites of Aspergillus candidus. II. The compound B (xanthoascin), a hepato-
and cardio-toxic xanthocillin analog. Chem. Pharm. Bull. 24, 2317–2321 (1976). 132
99. Ishida, M., Hamasaki, T. & Hatsuda, Y. The Structure of Two New Metabolites,
Emerin and Emericellin, from Aspergillus nidulans. Agricultural and Biological
Chemistry 39, 2181–2184 (1975).
100. Tsunakawa, M. et al. BU-4704, A NEW MEMBER OF THE XANTHOCILLIN
CLASS. J. Antibiot. 46, 687–688 (1993).
101. Zuck, K. M., Shipley, S. & Newman, D. J. Induced Production of N-Formyl
Alkaloids from Aspergillus fumigatus by Co-culture with Streptomyces peucetius.
J. Nat. Prod. 74, 1653–1657 (2011).
102. Morino, T., Nishimoto, M., Itou, N. & Nishikiori, T. NK372135S, NOVEL
ANTIFUNGAL AGENTS PRODUCED BY Neosartoria fischeri. J. Antibiot. 47,
1546–1548 (1994).
103. Takatsuki, A., Suzuki, S., Ando, K., Tamura, G. & Arima, K. NEW ANTIVIRAL
ANTIBIOTICS; XANTHOCILLIN X MONO- AND DIMETHYLETHER, AND
METHOXY- XANTHOCILLIN X DIMETHYLETHER. I. J. Antibiot. 21, 671–675
(1968).
104. Takatsuki, A., Yamaguchi, I., Tamura, G., Misato, T. & Arima, K.
CORRELATION BETWEEN THE ANTI-ANIMAL AND ANTI-PLANT-VIRUS
ACTIVITIES OF SEVERAL ANTIBIOTICS. J. Antibiot. 22, 442–445 (1969).
105. Itoh, J. et al. MK4588, A NEW ANTIBIOTIC RELATED TO XANTHOCILLIN.
J. Antibiot. 43, 456–461 (1990).
106. Zapf, S., HOßFELD, M., Anke, H., Velten, R. & Steglich, W. Darlucins A and B,
New Isocyanide Antibiotics from Sphaerellopsis filum (Darluca filum). J. Antibiot.
48, 36–41 (1995). 133
107. Vane, J. R. Inhibition of prostaglandin synthesis as a mechanism of action for
aspirin-like drugs. Nature New Biol. 231, 232–235 (1971).
108. Botting, R. M. Vane’s discovery of the mechanism of action of aspirin changed our
understanding of its clinical pharmacology. Pharmacological Reports 62, 518–525
(2010).
109. Boehm, J. C. et al. 1-Substituted 4-Aryl-5-pyridinylimidazoles: A New Class of
Cytokine Suppressive Drugs with Low 5-Lipoxygenase and Cyclooxygenase
Inhibitory Potency #. Journal of Medicinal Chemistry 39, 3929–3937 (1996).
110. Mechanism of the Van Leusen reaction. Henry Rzepa
https://www.ch.imperial.ac.uk/rzepa/blog/?p=10656 (2013).
111. Varghese, L. N., Defour, J.-P., Pecquet, C. & Constantinescu, S. N. The
Thrombopoietin Receptor: Structural Basis of Traffic and Activation by Ligand,
Mutations, Agonists, and Mutated Calreticulin. Front Endocrinol (Lausanne) 8,
(2017).
112. Singh, V. K., Kumar, N., Kalsan, M., Saini, A. & Chandra, R. A Novel Peptide
Thrombopoietin Mimetic Designing and Optimization Using Computational
Approach. Front Bioeng Biotechnol 4, 69 (2016).
113. Wang, Z. et al. Recombinant human thrombopoietin (rh-TPO) for the prevention of
severe thrombocytopenia induced by high-dose cytarabine: a prospective,
randomized, self-controlled study. Leuk. Lymphoma 1–8 (2018)
doi:10.1080/10428194.2018.1459605.
114. Bai, C., Xu, G., Zhao, Y., Han, S. & Shan, Y. [A multi-center clinical trial of
recombinant human thrombopoietin in the treatment of chemotherapy-induced 134
thrombocytopenia in patients with solid tumor]. Zhongguo Yi Xue Ke Xue Yuan Xue
Bao 26, 437–441 (2004).
115. Yamaguchi, T., Miyake, Y., Miyamura, A., Ishiwata, N. & Tatsuta, K. Structure-
activity Relationships of Xanthocillin Derivatives as Thrombopoietin Receptor
Agonist. The Journal of Antibiotics 59, 729–734 (2006).
116. Li, J. et al. Thrombocytopenia caused by the development of antibodies to
thrombopoietin. Blood 98, 3241–3248 (2001).
117. Sakai, R. et al. Xanthocillins as thrombopoietin mimetic small molecules.
Bioorganic & Medicinal Chemistry 13, 6388–6393 (2005).
118. Feuell, A. J. Toxic Factors of Mould Origin. Can Med Assoc J 94, 574–581 (1966).
119. McElhatton, P. R. Principles of teratogenicity. Current Obstetrics & Gynaecology 9,
163–169 (1999).
120. Takahashi, C., Sekita, S., Yoshihira, K. & Natori, S. The Structures of Toxic
Metabolites of Aspergillus candidus. II. The Compound B (Xanthoascin), a Hepato-
and Cardio-toxic Xanthocillin Analog. Chem. Pharm. Bull. 24, 2317–2321 (1976).
121. Ohtsubo, K. & Saito, M. Hepato-and cardiotoxicity of xanthoascin, a new
xanthocillin analogue produced by Aspergillus candidus. II. A preliminary electron
microscopic observation of the heart and lung with intranuclear myelin-like figures.
Ann Nutr Aliment 31, 771–780 (1977).
122. Ohtsubo, K., Horiuchi, T., Hatanaka, Y. & Saito, M. Hepato- and cardiotoxicity of
xanthoascin, a new metabolite of A. candidus Link, to mice. I. Blood chemistry and
histological changes in mice. Jpn J Exp Med 46, 277–287 (1976). 135
123. Kobayashi, J. et al. Brasilidine A, a New Cytotoxic Isonitrile-Containing Indole
Alkaloid from the Actinomycete Nocardia brasiliensis. J. Nat. Prod. 60, 719–720
(1997).
124. Naturally Occurring Isocyano/Isothiocyanato and Related Compounds |
SpringerLink. https://link.springer.com/chapter/10.1007/978-3-7091-6331-3_1.
125. Isaka, M., Boonkhao, B., Rachtawee, P. & Auncharoen, P. A xanthocillin-like
alkaloid from the insect pathogenic fungus Cordyceps brunnearubra BCC 1395. J.
Nat. Prod. 70, 656–658 (2007).
126. Mechanism of inhibiting proliferation by xanthocillin X dimethyl in tumor cells--《
Chinese Journal of New Drugs》2010年10期.
http://en.cnki.com.cn/Article_en/CJFDTOTAL-ZXYZ201010009.htm.
127. Zhao, Y. et al. SD118-Xanthocillin X (1), a Novel Marine Agent Extracted from
Penicillium commune, Induces Autophagy through the Inhibition of the MEK/ERK
Pathway. Marine Drugs 10, 1345–1359 (2012).
128. Nobuhara, M. et al. A Fungal Metabolite, Novel Isocyano Epoxide. Chem. Pharm.
Bull. 24, 832–834 (1976).
129. Pyke, T. R. & Dietz, A. U-21,963, a New Antibiotic: I. Discovery and Biological
Activity. Appl. Environ. Microbiol. 14, 506–510 (1966).
130. Tamura, A., Kotani, H. & Naruto, S. TRICHOVIRIDIN AND DERMADIN FROM
TRICHODERMA SP. TK-1. J. Antibiot. 28, 161–162 (1975).
131. Ollis, W. D. et al. The constitution of the antibiotic trichoviridin. Tetrahedron 36,
515–520 (1980). 136
132. Fujiwara, A. et al. Fermentation, Isolation and Characterization of Isonitrile
Antibiotics. Agricultural and Biological Chemistry 46, 1803–1809 (1982).
133. Baldwin, J. E. et al. Structure and synthesis of new cyclopentenyl isonitriles from
Trichoderma hamatum(Bon.) Bain. aggr. HLX 1379. Journal of the Chemical
Society, Chemical Communications 816 (1985) doi:10.1039/c39850000816.
134. Pratt, B. H., Sedgley, J. H., Heather, W. A. & Shepherd, C. J. Oospore Production in
Phytophthora Cinnamomi in the Presence of Trichoderma Koningii. Aust. Jnl. Of
Bio. Sci. 25, 861–864 (1972).
135. Reino, J. L., Guerrero, R. F., Hernández-Galán, R. & Collado, I. G. Secondary
metabolites from species of the biocontrol agent Type="Italic">Trichoderma
136. Faull, J. L., Graeme-Cook, K. A. & Pilkington, B. L. Production of an isonitrile
antibiotic by an UV-induced mutant of Trichoderma harzianum. Phytochemistry 36,
1273–1276 (1994).
137. Vinale, F. et al. Trichoderma Secondary Metabolites Active on Plants and Fungal
Pathogens. The Open Mycology Journal 8, (2014).
138. Potential risk factors associated with ill-thrift in buffalo calves (Bubalus bubalis)
raised at smallholder farms in Egypt. Journal of Advanced Research 6, 601–607
(2015).
139. Brewer, D. et al. Ovine ill-thrift in Nova Scotia. 9. Production of experimental
quantities of isocyanide metabolites of Trichoderma hamatum. Can. J. Microbiol.
28, 1252–1260 (1982). 137
140. Brewer, D. & Taylor, A. Ovine III-thrift in Nova Scotia. 7. Trichoderma hamatum
isolated from pasture soil. Mycopathologia 76, 167 (1981).
141. Fravel, D. R. Role of Antibiosis in the Biocontrol of Plant Diseases*. Annual
Review of Phytopathology 26, 75–91 (1988).
142. Liss, S. N., Brewer, D., Taylor, A. & Jones, G. A. Antibiotic activity of an
isocyanide metabolite of Trichoderma hamatum against rumen bacteria. Can. J.
Microbiol. 31, 767–772 (1985).
143. Feicht, A. & Taylor, A. Ovine ill-thrift in Nova Scotia. 8. Analysis of isocyanide
antibiotics produced by Trichoderma hamatum. Proceedings of the Nova Scotian
Institute of Science (1982).
144. Brewer, D., Calder, F. W., Jones, G. A., Tanguay, D. & Taylor, A. Effect of
Nickelous and Other Metal Ions on the Inhibition of Rumen Bacterial Metabolism
by 3-(3′-Isocyanocyclopent-2-Enylidene)Propionic Acid and Related Isocyanides.
Appl. Environ. Microbiol. 51, 138–142 (1986).
145. Cobalt deficiency in sheep and cattle. https://www.agric.wa.gov.au/livestock-
biosecurity/cobalt-deficiency-sheep-and-cattle.
146. Tiffany, M. E., Fellner, V. & Spears, J. W. Influence of cobalt concentration on
vitamin B12 production and fermentation of mixed ruminal microorganisms grown
in continuous culture flow-through fermentors. J Anim Sci 84, 635–640 (2006).
147. Sutherland, R. J., Cordes, D. O. & Carthew, G. C. Ovine white liver disease — an
hepatic dysfunction associated with vitamin B12 deficiency. New Zealand
Veterinary Journal 27, 227–232 (1979). 138
148. Vellema, P., van den Ingh, T. S. G. A. M. & Wouda, W. Pathological changes in
cobalt‐supplemented and non‐supplemented twin lambs in relation to blood
concentrations of methylmalonic acid and homocysteine. Veterinary Quarterly 21,
93–98 (1999).
149. Characterization of the kinetics and mechanisms of inhibition of drugs interacting
with the S. cerevisiae multidrug resistance pumps Pdr5p and Snq2p. Biochimica et
Biophysica Acta (BBA) - Biomembranes 1788, 717–723 (2009).
150. Rutledge, R. M., Esser, L., Ma, J. & Xia, D. Toward understanding the mechanism
of action of the yeast multidrug resistance transporter Pdr5p: a molecular modeling
study. J. Struct. Biol. 173, 333–344 (2011).
151. A new function of isonitrile as an inhibitor of the Pdr5p multidrug ABC transporter
in Saccharomyces cerevisiae. Biochemical and Biophysical Research
Communications 330, 622–628 (2005).
152. Inhibition of fatty acid synthetases by the antibiotic cerulenin. Biochemical and
Biophysical Research Communications 48, 649–656 (1972).
153. Imada, C. et al. Isolation and characterization of tyrosinase inhibitor-producing
microorganisms from marine environment. Fisheries science 67, 1151–1156 (2001).
154. Fernandes, M. S. & Kerkar, S. Microorganisms as a source of tyrosinase inhibitors:
a review. Ann Microbiol 67, 343–358 (2017).
155. Chang, T.-S. & Chang, T.-S. An Updated Review of Tyrosinase Inhibitors.
International Journal of Molecular Sciences 10, 2440–2475 (2009).
156. Chang, S. T., Shepherd, C. J. & Pratt, B. H. Control of sexuality in Phytophthora
cinnamomi. Aust. J. Bot. 22, 669–679 (1974). 139
157. Brasier, C. M. Induction of Sexual Reproduction in Single A2 Isolates of
Phytophthora Species by Trichoderma viride. Nature New Biology 231, 283 (1971).
158. Brasier, C. M. Stimulation of Sex Organ Formation in Phytophthora by Antagonistic
Species of Trichoderma. New Phytologist 74, 195–198 (1975).
159. Martins, N., Ferreira, I. C. F. R., Barros, L., Silva, S. & Henriques, M. Candidiasis:
Predisposing Factors, Prevention, Diagnosis and Alternative Treatment.
Mycopathologia 177, 223–240 (2014).
160. Turner, J. R., Butler, T. F., Gordee, R. S. & Thakkar, A. L. A32390A, A NEW
BIOLOGICALLY ACTIVE METABOLITE. J. Antibiot. 31, 33–37 (1978).
161. Marconi, G. G. et al. A32390A, A NEW BIOLOGICALLY ACTIVE
METABOLITE. J. Antibiot. 31, 27–32 (1978).
162. Rush, R. A. & Geffen, L. B. Dopamine β-Hydroxylase in Health and Disease. CRC
Critical Reviews in Clinical Laboratory Sciences 12, 241–277 (1980).
163. KAUFMAN, S., BRIDGERS, W. F. & BARON, J. The Mechanism of Action of
Dopamine β-Hydroxylase. in Oxidation of Organic Compounds vol. 77 172–176
(AMERICAN CHEMICAL SOCIETY, 1968).
164. Robertson, D. et al. Dopamine beta-hydroxylase deficiency. A genetic disorder of
cardiovascular regulation. Hypertension 18, 1–8 (1991).
165. Cross, A. J. et al. Reduced dopamine-beta-hydroxylase activity in Alzheimer’s
disease. Br Med J (Clin Res Ed) 282, 93–94 (1981).
166. Association of the dopamine beta hydroxylase gene with attention deficit
hyperactivity disorder: Genetic analysis of the Milwaukee longitudinal study - 140
Smith - 2003 - American Journal of Medical Genetics Part B: Neuropsychiatric
Genetics - Wiley Online Library.
167. Plasma dopamine beta-hydroxylase activity in psychotic and non-psychotic post-
traumatic stress disorder. Psychiatry Research 77, 175–181 (1998).
168. Decreased dopamine beta-hydroxylase activity in unipolar geriatric delusional
depression. Biological Psychiatry 45, 448–452 (1999).
169. Boeck, L. D., Hoehn, M. M., Sands, T. H. & Wetzel, R. W. A32390A, A NEW
BIOLOGICALLY ACTIVE METABOLITE. J. Antibiot. 31, 19–26 (1978).
170. New inhibitors of dopamine β-hydroxylase. Advances in Enzyme Regulation 15,
267–281 (1977).
171. Palfreyman, M. Enzymes as Targets for Drug Design. (Elsevier, 2012).
172. Ciegler, A. & Lindenfelser, L. A. An antibiotic complex from Type="Italic">Alternaria brassicicola
(1969).
173. Gloer, J. B., Poch, G. K., Short, D. M. & McCloskey, D. V. Structure of
brassicicolin A: a novel isocyanide antibiotic from the phylloplane fungus
Alternaria brassicicola. J. Org. Chem. 53, 3758–3761 (1988).
174. The phytopathogenic fungus Alternaria brassicicola: Phytotoxin production and
phytoalexin elicitation. Phytochemistry 70, 394–402 (2009).
175. Pedras, M. S. C. & Park, M. R. Metabolite diversity in the plant pathogen Alternaria
brassicicola: factors affecting production of brassicicolin A, depudecin,
phomapyrone A and other metabolites. Mycologia 107, 1138–1150 (2015). 141
176. Evans, J. R., Napier, E. J. & Yates, P. ISOLATION OF A NEW ANTIBIOTIC
FROM A SPECIES OF PSEUDOMONAS. J. Antibiot. 29, 850–852 (1976).
177. Marquez, J. A. et al. THE HAZIMICINS, A NEW CLASS OF ANTIBIOTICS. J.
Antibiot. 36, 1101–1108 (1983).
178. Wright, J. J. K. et al. X-Ray crystal structure determination and synthesis of the new
isonitrile-containing antibiotics, hazimycin factors 5 and 6. J. Chem. Soc., Chem.
Commun. 0, 1188–1190 (1982).
179. Parker, W. L. et al. AEROCYANIDIN, A NEW ANTIBIOTIC PRODUCED BY
CHROMOBACTERIUM VIOLACEUM. J. Antibiot. 41, 454–460 (1988).
180. Singh, P. D. et al. AEROCAVIN, A NEW ANTIBIOTIC PRODUCED BY
CHROMOBACTERIUM VIOLACEUM. J. Antibiot. 41, 446–453 (1988).
181. J. Garson, M. & S. Simpson, J. Marine isocyanides and related natural products –
structure, biosynthesis and ecology. Natural Product Reports 21, 164–179 (2004).
182. Chang, C. W. J. & Scheuer, P. J. Marine isocyano compounds. in Marine Natural
Products — Diversity and Biosynthesis (ed. Scheuer, P. J.) vol. 167 33–75
(Springer-Verlag, 1993).
183. Moore, R. E. et al. Hapalindoles, antibacterial and antimycotic alkaloids from the
cyanophyte Hapalosiphon fontinalis. The Journal of Organic Chemistry 52, 1036–
1043 (1987).
184. Moore, R. E., Cheuk, C. & Patterson, G. M. L. Hapalindoles: new alkaloids from
the blue-green alga Hapalosiphon fontinalis. Journal of the American Chemical
Society 106, 6456–6457 (1984). 142
185. Bhat, V., Dave, A., MacKay, J. A. & Rawal, V. H. The Chemistry of Hapalindoles,
Fischerindoles, Ambiguines, and Welwitindolinones. Alkaloids Chem Biol 73, 65–
160 (2014).
186. Borowitzka, M. A. Microalgae as sources of pharmaceuticals and other biologically
active compounds. J Appl Phycol 7, 3–15 (1995).
187. Kim, H. et al. Indole alkaloids from two cultured cyanobacteria, Westiellopsis sp.
and Fischerella muscicola. Bioorganic & Medicinal Chemistry 20, 5290–5295
(2012).
188. Klein, D., Daloze, D., Braekman, J. C., Hoffmann, L. & Demoulin, V. New
Hapalindoles from the Cyanophyte Hapalosiphon laingii. J. Nat. Prod. 58, 1781–
1785 (1995).
189. Bonjouklian, R., Moore, R. E., Patterson, G. M. L. & Smitka, T. A. Preparation of
hapalindole-related alkaloids from blue-green algae. (1994).
190. Smitka, T. A. et al. Ambiguine isonitriles, fungicidal hapalindole-type alkaloids
from three genera of blue-green algae belonging to the Stigonemataceae. The
Journal of Organic Chemistry 57, 857–861 (1992).
191. Park, A., Moore, R. E. & Patterson, G. M. L. Fischerindole L, a new isonitrile from
the terrestrial blue-green alga fischerella muscicola. Tetrahedron Letters 33, 3257–
3260 (1992).
192. Asthana, R. K. et al. Identification of an antimicrobial entity from the
cyanobacterium Fischerella sp. isolated from bark of Azadirachta indica (Neem)
tree. J Appl Phycol 18, 33–39 (2006). 143
193. Becher, P. G., Keller, S., Jung, G., Süssmuth, R. D. & Jüttner, F. Insecticidal
activity of 12-epi-hapalindole J isonitrile. Phytochemistry 68, 2493–2497 (2007).
194. Sanevas, N., Sunohara, Y. & Matsumoto, H. Crude extract of the cyanobacterium,
Hapalosiphon sp., causes a cessation of root elongation and cell division in several
plant species. Weed Biology and Management 6, 25–29 (2006).
195. Sanevas, N., Sunohara, Y. & Matsumoto, H. Characterization of reactive oxygen
species-involved oxidative damage in Hapalosiphon species crude extract-treated
wheat and onion roots. Weed Biology and Management 7, 172–177 (2007).
196. Isolation of ambiguine D isonitrile from Hapalosiphon sp. and characterization of its
phytotoxic activity | SpringerLink.
https://link.springer.com/article/10.1007%2Fs10725-012-9700-8.
197. Ready, J. M. et al. A Mild and Efficient Synthesis of Oxindoles: Progress Towards
the Synthesis of Welwitindolinone A Isonitrile. Angewandte Chemie International
Edition 43, 1270–1272 (2004).
198. Smith, C. D., Zilfou, J. T., Stratmann, K., Patterson, G. M. & Moore, R. E.
Welwitindolinone analogues that reverse P-glycoprotein-mediated multiple drug
resistance. Mol Pharmacol 47, 241–247 (1995).
199. Zhang, X. & Smith, C. D. Microtubule effects of welwistatin, a cyanobacterial
indolinone that circumvents multiple drug resistance. Molecular Pharmacology 49,
288–294 (1996).
200. Doan, N. T., Stewart, P. R. & Smith, G. D. Inhibition of bacterial RNA polymerase
by the cyanobacterial metabolites 12-epi-hapalindole E isonitrile and calothrixin A.
FEMS Microbiology Letters 196, 135–139 (2001). 144
201. Kerry, L. E. et al. Selective inhibition of RNA polymerase I transcription as a
potential approach to treat African trypanosomiasis. PLOS Neglected Tropical
Diseases 11, e0005432 (2017).
202. Bywater, M. J. et al. Inhibition of RNA Polymerase I as a Therapeutic Strategy to
Promote Cancer-Specific Activation of p53. Cancer Cell 22, 51–65 (2012).
203. Hannan, R. D., Drygin, D. & Pearson, R. B. Targeting RNA polymerase I
transcription and the nucleolus for cancer therapy. Expert Opinion on Therapeutic
Targets 17, 873–878 (2013).
204. Hein, N., Hannan, K. M., D’Rozario, J. & Hannan, R. Inhibition of Pol I
Transcription a New Chance in the Fight Against Cancer. Technol Cancer Res Treat
16, 736–739 (2017).
205. Lee, H. et al. RNA Polymerase I Inhibition with CX-5461 As a Novel Strategy to
Target MYC in Multiple Myeloma. Blood 126, 3010–3010 (2015).
206. Schwartz, R. E., Hirsch, C. F., Sigmund, J. M. & Pettibone, D. J. Hapalindolinone
compounds as vassopressin antagonists. (1989).
207. Schwartz, R. E., Hirsch, C. F., Springer, J. P., Pettibone, D. J. & Zink, D. L.
Unusual cyclopropane-containing hapalindolinones from a cultured
cyanobacterium. J. Org. Chem. 52, 3704–3706 (1987).
208. Richter, J. M. et al. Enantiospecific Total Synthesis of the Hapalindoles,
Fischerindoles, and Welwitindolinones via a Redox Economic Approach. Journal of
the American Chemical Society 130, 17938–17954 (2008). 145
209. Ittiamornkul, K. et al. Promiscuous indolyl vinyl isonitrile synthases in the
biogenesis and diversification of hapalindole-type alkaloids. Chemical Science 6,
6836–6840 (2015).
210. Burreson, B. J. & Scheuer, P. J. Isolation of a diterpenoid isonitrile from a marine
sponge. Journal of the Chemical Society, Chemical Communications 1035 (1974)
doi:10.1039/c39740001035.
211. Patra, A. et al. An unprecedented triisocyano diterpenoid antibiotic from a sponge.
Journal of the American Chemical Society 106, 7981–7983 (1984).
212. Chang, C. W. J. et al. Kalihinol-A, a highly functionalized diisocyano diterpenoid
antibiotic from a sponge. Journal of the American Chemical Society 106, 4644–
4646 (1984).
213. Trimurtulu, G. & Faulkner, D. J. Six New Diterpene Isonitriles from the Sponge
Acanthella cavernosa. J. Nat. Prod. 57, 501–506 (1994).
214. Fusetani, N. et al. Kalihinene and isokalihinol B, cytotoxic diterpene isonitriles
from the marine sponge Acanthella klethra. Tetrahedron Letters 31, 3599–3602
(1990).
215. Alvi, K. A., Tenenbaum, L. & Crews, P. Anthelmintic Polyfunctional Nitrogen-
Containing Terpenoids from Marine Sponges. J. Nat. Prod. 54, 71–78 (1991).
216. Mendola, D. Aquaculture of three phyla of marine invertebrates to yield bioactive
metabolites: process developments and economics. Biomol. Eng. 20, 441–458
(2003).
217. Miyaoka, H. et al. Antimalarial activity of kalihinol A and new relative diterpenoids
from the Okinawan sponge, Acanthella sp. Tetrahedron 54, 13467–13474 (1998). 146
218. Wright, A. D. et al. Antimalarial Activity: The Search for Marine-Derived Natural
Products with Selective Antimalarial Activity. J. Nat. Prod. 59, 710–716 (1996).
219. Okino, T., Yoshimura, E., Hirota, H. & Fusetani, N. Antifouling kalihinenes from
the marine sponge Acanthella cavernosa. Tetrahedron Letters 36, 8637–8640
(1995).
220. Okino, T., Yoshimura, E., Hirota, H. & Fusetani, N. New Antifouling Kalihipyrans
from the Marine Sponge Acanthella cavernosa. J. Nat. Prod. 59, 1081–1083 (1996).
221. Wratten, S. J., Faulkner, D. J., Hirotsu, K. & Clardy, J. Diterpenoid isocyanides
from the marine sponge hymeniacidon amphilecta. Tetrahedron Letters 19, 4345–
4348 (1978).
222. König, G. M., Wright, A. D. & Angerhofer, C. K. Novel Potent Antimalarial
Diterpene Isocyanates, Isothiocyanates, and Isonitriles from the Tropical Marine
Sponge Cymbastela hooperi. J. Org. Chem. 61, 3259–3267 (1996).
223. Kazlauskas, R., Murphy, P. T., Wells, R. J. & Blount, J. F. New diterpene
isocyanides from a sponge. Tetrahedron Letters 21, 315–318 (1980).
224. Molinski, T. F., Faulkner, D. J., Van Duyne, G. D. & Clardy, J. Three new diterpene
isonitriles from a Palauan sponge of the genus Halichondria. The Journal of
Organic Chemistry 52, 3334–3337 (1987).
225. Sharma, H. A. et al. Two New Diterpene Isocyanides from a Sponge of the Family
Adocidae. Tetrahedron Letters 33, 1593–1596 (1992).
226. Fookes, C. J. R., Garson, M. J., MacLeod, J. K., Skelton, B. W. & White, A. H.
Biosynthesis of diisocyanoadociane, a novel diterpene from the marine sponge 147
Amphimedon sp. Crystal structure of a monoamide derivative. J. Chem. Soc.,
Perkin Trans. 1 0, 1003–1011 (1988).
227. Schwarz, O., Brun, R., Bats, J. W. & Schmalz, H.-G. Synthesis and biological
evaluation of new antimalarial isonitriles related to marine diterpenoids.
Tetrahedron Letters 43, 1009–1013 (2002).
228. Singh, C., Srivastav, N. C. & Puri, S. K. In vivo active antimalarial isonitriles.
Bioorganic & Medicinal Chemistry Letters 12, 2277–2279 (2002).
229. Tekwani, B. L. & Walker, L. A. Targeting the hemozoin synthesis pathway for new
antimalarial drug discovery: technologies for in vitro beta-hematin formation assay.
Comb. Chem. High Throughput Screen. 8, 63–79 (2005).
230. Kumar, S., Guha, M., Choubey, V., Maity, P. & Bandyopadhyay, U. Antimalarial
drugs inhibiting hemozoin (beta-hematin) formation: a mechanistic update. Life Sci.
80, 813–828 (2007).
231. Wright, C. W. et al. Synthesis and Evaluation of Cryptolepine Analogues for Their
Potential as New Antimalarial Agents. J. Med. Chem. 44, 3187–3194 (2001).
232. Fattorusso, E., Magno, S., Mayol, L., Santacroce, C. & Sica, D. Isolation and
structure of axisonitrile-2: A new sesquiterpenoid isonitrile from the sponge
Axinella cannabina. Tetrahedron 30, 3911–3913 (1974).
233. Minale, L., Riccio, R. & Sodano, G. Acanthellin-1, an unique isonitrile
sesquiterpene from the sponge Acanthella acuta. Tetrahedron 30, 1341–1343
(1974). 148
234. Ciminiello, P., Fattorusso, E., Magno, S. & Mayol, L. New nitrogenous
sesquiterpenes from the marine sponge Axinella cannabina. The Journal of Organic
Chemistry 49, 3949–3951 (1984).
235. Thompson, J. E., Walker, R. P., Wratten, S. J. & Faulkner, D. J. A chemical defense
mechanism for the nudibranch cadlina luteomarginata. Tetrahedron 38, 1865–1873
(1982).
236. Di Blasio, B. et al. Axisonitrile-3, axisothiocyanate-3 and axamide-3.
Sesquiterpenes with a novel spiro[4,5]decane skeleton from the sponge Axinella
cannabina. Tetrahedron 32, 473–478 (1976).
237. Mayol, L., Piccialli, V. & Sica, D. Nitrogenous sesquiterpenes from the marine
sponge acanthella acuta: three new isocyanide-isothiocyanate pairs. Tetrahedron 43,
5381–5388 (1987).
238. Angerhofer, C. K., Pezzuto, J. M., König, G. M., Wright, A. D. & Sticher, O.
Antimalarial Activity of Sesquiterpenes from the Marine Sponge Acanthella klethra.
Journal of Natural Products 55, 1787–1789 (1992).
239. He, H. Y., Salva, J., Catalos, R. F. & Faulkner, D. J. Sesquiterpene thiocyanates and
isothiocyanates from Axinyssa aplysinoides. The Journal of Organic Chemistry 57,
3191–3194 (1992).
240. Fusetani, N. et al. Two sesquiterpene isocyanides and a sesquiterpene thiocyanate
from the marine sponge Acanthella cf. cavernosa and the Nudibranch Phyllidia
ocellata. Tetrahedron Letters 33, 6823–6826 (1992). 149
241. Hirota, H., Tomono, Y. & Fusetani, N. Terpenoids with antifouling activity against
barnacle larvae from the marine sponge Acanthella cavernosa. Tetrahedron 52,
2359–2368 (1996).
242. Okino, T., Yoshimura, E., Hirota, H. & Fusetani, N. New antifouling sesquiterpenes
from four nudibranchs of the family Phyllidiidae. Tetrahedron 52, 9447–9454
(1996).
243. Nakamura, H., Kobayashi, J., Ohizumi, Y., Mitsubishi-Kasei & Hirata, Y. Novel
bisabolene-type sesquiterpenoids with a conjugated diene isolated from the
okinawan sea sponge theonella cf. swinhoei. Tetrahedron Letters 25, 5401–5404
(1984).
244. Burreson, B. J., Scheuer, P. J., Finer, J. & Clardy, J. 9-Isocyanopupukeanane, a
marine invertebrate allomone with a new sesquiterpene skeleton. Journal of the
American Chemical Society 97, 4763–4764 (1975).
245. Hagadone, M. R., Burreson, B. J., Scheuer, P. J., Finer, J. S. & Clardy, J. Defense
Allomones of the Nudibranch Phyllidia varicosa Lamarck 1801. Helvetica Chimica
Acta 62, 2484–2494 (1979).
246. Amade, Ph., Pesando, D. & Chevolot, L. Antimicrobial activities of marine sponges
from French Polynesia and Brittany. Mar. Biol. 70, 223–228 (1982).
247. Burreson, B. J., Christophersen, C. & Scheuer, P. J. Co-occurrence of two terpenoid
isocyanide-formamide pairs in a marine sponge (Halichondria sp.). Tetrahedron 31,
2015–2018 (1975).
248. Gijsen, H. J. M., Wijnberg, J. B. P. A. & De Groot, Ae. Structure, Occurrence,
Biosynthesis, Biological Activity, Synthesis, and Chemistry of Aromadendrane 150
Sesquiterpenoids. in Fortschritte der Chemie organischer Naturstoffe / Progress in
the Chemistry of Organic Natural Products (eds. Barrera, J. B. et al.) 149–193
(Springer Vienna, 1995). doi:10.1007/978-3-7091-9337-2_3.
249. Capon, R. J. & Macleod, J. K. New Isothiocyanate Sesquiterpenes From the
Australian Marine Sponge Acanthella pulcherrima. Aust. J. Chem. 41, 979–983
(1988).
250. Thompson, J. E., Walker, R. P. & Faulkner, D. J. Screening and bioassays for
biologically-active substances from forty marine sponge species from San Diego,
California, USA. Mar. Biol. 88, 11–21 (1985).
251. Braekman, J. C., Daloze, D., Moussiaux, B., Stoller, C. & Deneubourg, F. Sponge
secondary metabolites: new results. Pure and Applied Chemistry 61, 509–512
(1989).
252. Braekman, J. C., Daloze, D., Deneubourg, F., Huysecom, J. & Vandevyver, G. I-
Isocyanoaromadendrane, A New Isonitrile Sesquiterpene from the Sponge
Acanthella Acuta. Bulletin des Sociétés Chimiques Belges 96, 539–543 (1987).
253. Goda, M. et al. Isonitrile Hydratase from Pseudomonas putidaN19–2 CLONING,
SEQUENCING, GENE EXPRESSION, AND IDENTIFICATION OF ITS
ACTIVE AMINO ACID RESIDUE. J. Biol. Chem. 277, 45860–45865 (2002).
254. Sato, H., Hashimoto, Y., Fukatsu, H. & Kobayashi, M. Novel Isonitrile Hydratase
Involved in Isonitrile Metabolism. J. Biol. Chem. 285, 34793–34802 (2010).
255. Pineda, J. R. E. T., Antoniou, D. & Schwartz, S. D. Slow Conformational Motions
That Favor Sub-picosecond Motions Important for Catalysis. J. Phys. Chem. B 114,
15985–15990 (2010). 151
256. Hay, S. & Scrutton, N. S. Good vibrations in enzyme-catalysed reactions. Nature
Chemistry 4, 161–168 (2012).
257. Olsson, M. H. M., Parson, W. W. & Warshel, A. Dynamical Contributions to
Enzyme Catalysis: Critical Tests of A Popular Hypothesis. Chem. Rev. 106, 1737–
1756 (2006).
258. Mulholland, A. J., Roitberg, A. E. & Tuñón, I. Enzyme dynamics and catalysis in
the mechanism of DNA polymerase. Theor Chem Acc 131, 1286 (2012).
259. Agarwal, P. K., Billeter, S. R., Rajagopalan, P. T. R., Benkovic, S. J. & Hammes-
Schiffer, S. Network of coupled promoting motions in enzyme catalysis. PNAS 99,
2794–2799 (2002).
260. Glowacki, D. R., Harvey, J. N. & Mulholland, A. J. Protein dynamics and enzyme
catalysis: the ghost in the machine? Biochem Soc Trans 40, 515–521 (2012).
261. Benkovic, S. J. & Hammes-Schiffer, S. A Perspective on Enzyme Catalysis. Science
301, 1196–1202 (2003).
262. Nashine, V. C., Hammes-Schiffer, S. & Benkovic, S. J. Coupled motions in enzyme
catalysis. Current Opinion in Chemical Biology 14, 644–651 (2010).
263. Antoniou, D., Basner, J., Núñez, S. & Schwartz, S. D. Computational and
Theoretical Methods to Explore the Relation between Enzyme Dynamics and
Catalysis. Chem. Rev. 106, 3170–3187 (2006).
264. Kamerlin, S. C. L. & Warshel, A. At the dawn of the 21st century: Is dynamics the
missing link for understanding enzyme catalysis? Proteins: Structure, Function, and
Bioinformatics 78, 1339–1375 (2010). 152
265. DeCarlo, L., Gowda, A. S. P., Suo, Z. & Spratt, T. E. Formation of Purine−Purine
Mispairs by Sulfolobus solfataricus DNA Polymerase IV. Biochemistry 47, 8157–
8164 (2008).
266. Guengerich, F. P. Interactions of Carcinogen-Bound DNA with Individual DNA
Polymerases. Chem. Rev. 106, 420–452 (2006).
267. Mizrahi, V., Henrie, R. N., Marlier, J. F., Johnson, K. A. & Benkovic, S. J. Rate-
limiting steps in the DNA polymerase I reaction pathway. Biochemistry 24, 4010–
4018 (1985).
268. Wolf-Watz, M. et al. Linkage between dynamics and catalysis in a thermophilic-
mesophilic enzyme pair. Nature Structural & Molecular Biology 11, 945–949
(2004).
269. Hammes, G. G. Multiple Conformational Changes in Enzyme Catalysis †.
Biochemistry 41, 8221–8228 (2002).
270. Watt, E. D., Shimada, H., Kovrigin, E. L. & Loria, J. P. The mechanism of rate-
limiting motions in enzyme function. Proceedings of the National Academy of
Sciences 104, 11981–11986 (2007).
271. Active site dynamics of ribonuclease | PNAS.
https://www.pnas.org/content/82/24/8458.short.
272. Messmore, J. M., Fuchs, D. N. & Raines, R. T. Ribonuclease A: Revealing
Structure–Function Relationships with Semisynthesis. J Am Chem Soc 117, 8057–
8060 (1995).
273. Park, C. & Raines, R. T. Catalysis by Ribonuclease A Is Limited by the Rate of
Substrate Association. Biochemistry 42, 3509–3518 (2003). 153
274. Relaxation Spectra of Ribonuclease. I. The Interaction of Ribonuclease with
Cytidine 3′-Phosphate | Journal of the American Chemical Society.
https://pubs.acs.org/doi/pdf/10.1021/ja01070a008.
275. Beach, H., Cole, R., Gill, M. L. & Loria, J. P. Conservation of μs−ms Enzyme
Motions in the Apo- and Substrate-Mimicked State. J. Am. Chem. Soc. 127, 9167–
9176 (2005).
276. Enzyme Dynamics During Catalysis | Science.
https://science.sciencemag.org/content/295/5559/1520?casa_token=iYJKPMEpYlc
AAAAA:WFdcvQI9dPuZSSFH0UkkiXgBf4vYZV8UoU0lkzxL8TaC6xseu1dwYX
r63vLILZ2alSxOwbl_Ay63ous.
277. Intrinsic dynamics of an enzyme underlies catalysis | Nature.
https://www.nature.com/articles/nature04105.
278. Boehr, D. D., McElheny, D., Dyson, H. J. & Wright, P. E. The Dynamic Energy
Landscape of Dihydrofolate Reductase Catalysis. Science 313, 1638–1642 (2006).
279. Cole, R. & Loria, J. P. Evidence for Flexibility in the Function of Ribonuclease A.
Biochemistry 41, 6072–6081 (2002).
280. Kartha, G., Bello, J. & Harker, D. Tertiary Structure of Ribonuclease. Nature 213,
862–865 (1967).
281. Kovrigin, E. L., Cole, R. & Loria, J. P. Temperature Dependence of the Backbone
Dynamics of Ribonuclease A in the Ground State and Bound to the Inhibitor 5‘-
Phosphothymidine (3‘-5‘)Pyrophosphate Adenosine 3‘-Phosphate. Biochemistry 42,
5279–5291 (2003). 154
282. Zegers, I. et al. The structures of rnase a complexed with 3′-CMP and d(CpA):
Active site conformation and conserved water molecules. Protein Science 3, 2322–
2339 (1994).
283. Dubins, D. N., Filfil, R., Macgregor, R. B. & Chalikian, T. V. Role of Water in
Protein−Ligand Interactions: Volumetric Characterization of the Binding of 2‘-
CMP and 3‘-CMP to Ribonuclease A. J. Phys. Chem. B 104, 390–401 (2000).
284. Structure of phosphate-free ribonuclease A refined at 1.26 .ANG. | Biochemistry.
https://pubs.acs.org/doi/abs/10.1021/bi00408a010.
285. Pauling, L. Nature of Forces between Large Molecules of Biological Interest*.
Nature 161, 707–709 (1948).
286. Jencks, W. P. Binding Energy, Specificity, and Enzymic Catalysis: The Circe
Effect. in Advances in Enzymology and Related Areas of Molecular Biology 219–
410 (John Wiley & Sons, Ltd, 2006). doi:10.1002/9780470122884.ch4.
287. Menard, R. et al. Contribution of the glutamine 19 side chain to transition-state
stabilization in the oxyanion hole of papain. Biochemistry 30, 8924–8928 (1991).
288. Schramm, V. L. Enzymatic transition states and transition state analogues. Current
Opinion in Structural Biology 15, 604–613 (2005).
289. Structural Biochemistry/Enzyme/Transition state - Wikibooks, open books for an
open world.
https://en.m.wikibooks.org/wiki/Structural_Biochemistry/Enzyme/Transition_state.
290. Tesmer, J. J. G., Berman, D. M., Gilman, A. G. & Sprang, S. R. Structure of RGS4
Bound to AlF4−-Activated Giα1: Stabilization of the Transition State for GTP
Hydrolysis. Cell 89, 251–261 (1997). 155
291. Štrajbl, M., Shurki, A., Kato, M. & Warshel, A. Apparent NAC Effect in
Chorismate Mutase Reflects Electrostatic Transition State Stabilization. J. Am.
Chem. Soc. 125, 10228–10237 (2003).
292. Rupert, P. B., Massey, A. P., Sigurdsson, S. T. & Ferré-D’Amaré, A. R. Transition
State Stabilization by a Catalytic RNA. Science 298, 1421–1424 (2002).
293. Leatherbarrow, R. J., Fersht, A. R. & Winter, G. Transition-state stabilization in the
mechanism of tyrosyl-tRNA synthetase revealed by protein engineering. PNAS 82,
7840–7844 (1985).
294. Andrews, P. R., Smith, G. D. & Young, I. G. Transition-state stabilization and
enzymic catalysis. Kinetic and molecular orbital studies of the rearrangement of
chorismate to prephenate. Biochemistry 12, 3492–3498 (1973).
295. Robertus, J. D., Kraut, J., Alden, R. A. & Birktoft, J. J. Subtilisin. Stereochemical
mechanism involving transition-state stabilization. Biochemistry 11, 4293–4303
(1972).
296. Stewart, J. D. & Benkovic, S. J. Transition-state stabilization as a measure of the
efficiency of antibody catalysis. Nature 375, 388–391 (1995).
297. Hall, A., Parsonage, D., Poole, L. B. & Karplus, P. A. Structural Evidence that
Peroxiredoxin Catalytic Power Is Based on Transition-State Stabilization. Journal of
Molecular Biology 402, 194–209 (2010).
298. The .alpha. effect: on the origin of transition-state stabilization.
https://pubs.acs.org/doi/pdf/10.1021/jo00139a033 doi:10.1021/jo00139a033.
299. Warshel, A., Štrajbl, M., Villà, J. & Florián, J. Remarkable Rate Enhancement of
Orotidine 5‘-Monophosphate Decarboxylase Is Due to Transition-State Stabilization 156
Rather Than to Ground-State Destabilization. Biochemistry 39, 14728–14738
(2000).
300. Alhambra, C., Wu, L., Zhang, Z.-Y. & Gao, J. Walden-Inversion-Enforced
Transition-State Stabilization in a Protein Tyrosine Phosphatase. J. Am. Chem. Soc.
120, 3858–3866 (1998).
301. Nadanaciva, S., Weber, J., Wilke-Mounts, S. & Senior, A. E. Importance of F1-
ATPase Residue α-Arg-376 for Catalytic Transition State Stabilization.
Biochemistry 38, 15493–15499 (1999).
302. Electrostatic Basis for Enzyme Catalysis | Chemical Reviews.
https://pubs.acs.org/doi/abs/10.1021/cr0503106.
303. Hansson, T., Nordlund, P. & Åqvist, J. Energetics of nucleophile activation in a
protein tyrosine phosphatase11Edited by A. Fersht. Journal of Molecular Biology
265, 118–127 (1997).
304. Warshel, A., Sharma, P. K., Chu, Z. T. & Åqvist, J. Electrostatic Contributions to
Binding of Transition State Analogues Can Be Very Different from the
Corresponding Contributions to Catalysis: Phenolates Binding to the Oxyanion Hole
of Ketosteroid Isomerase †. Biochemistry 46, 1466–1476 (2007).
305. Schowen, K. B., Limbach, H.-H., Denisov, G. S. & Schowen, R. L. Hydrogen bonds
and proton transfer in general-catalytic transition-state stabilization in enzyme
catalysis. Biochimica et Biophysica Acta (BBA) - Bioenergetics 1458, 43–62 (2000).
306. Breslow, R. & Chapman, W. H. On the mechanism of action of ribonuclease A:
relevance of enzymatic studies with a p-nitrophenylphosphate ester and a
thiophosphate ester. PNAS 93, 10018–10021 (1996). 157
307. Schwartz, S. D. & Schramm, V. L. Enzymatic transition states and dynamic motion
in barrier crossing. Nature Chemical Biology 5, 551–558 (2009).
308. Bystroff, C. & Kraut, J. Crystal structure of unliganded Escherichia coli
dihydrofolate reductase. Ligand-induced conformational changes and cooperativity
in binding. Biochemistry 30, 2227–2239 (1991).
309. Volz, K. W. et al. Crystal structure of avian dihydrofolate reductase containing
phenyltriazine and NADPH. J. Biol. Chem. 257, 2528–2536 (1982).
310. Li, L., Wright, P. E., Benkovic, S. J. & Falzone, C. J. Functional role of a mobile
loop of Escherichia coli dihydrofolate reductase in transition-state stabilization.
Biochemistry 31, 7826–7833 (1992).
311. Penner, M. H. & Frieden, C. Kinetic analysis of the mechanism of Escherichia coli
dihydrofolate reductase. J. Biol. Chem. 262, 15908–15914 (1987).
312. Bhabha, G. et al. A Dynamic Knockout Reveals That Conformational Fluctuations
Influence the Chemical Step of Enzyme Catalysis. Science 332, 234–238 (2011).
313. Schnell, J. R., Dyson, H. J. & Wright, P. E. Structure, Dynamics, and Catalytic
Function of Dihydrofolate Reductase. Annual Review of Biophysics and
Biomolecular Structure 33, 119–140 (2004).
314. Epstein, D. M., Benkovic, S. J. & Wright, P. E. Dynamics of the Dihydrofolate
Reductase-Folate Complex: Catalytic Sites and Regions Known To Undergo
Conformational Change Exhibit Diverse Dynamical Features. Biochemistry 34,
11037–11048 (1995). 158
315. Cameron, C. E. & Benkovic, S. J. Evidence for a Functional Role of the Dynamics
of Glycine-121 of Escherichia coli Dihydrofolate Reductase Obtained from Kinetic
Analysis of a Site-Directed Mutant. Biochemistry 36, 15792–15800 (1997).
316. Schramm, V. L. Enzymatic Transition States, Transition-State Analogs, Dynamics,
Thermodynamics, and Lifetimes. Annual Review of Biochemistry 80, 703–732
(2011).
317. Adamczyk, A. J., Cao, J., Kamerlin, S. C. L. & Warshel, A. Catalysis by
dihydrofolate reductase and other enzymes arises from electrostatic preorganization,
not conformational motions. Proceedings of the National Academy of Sciences 108,
14115–14120 (2011).
318. Olsson, M. H. M., Mavri, J. & Warshel, A. Transition state theory can be used in
studies of enzyme catalysis: lessons from simulations of tunnelling and dynamical
effects in lipoxygenase and other systems. Philosophical Transactions of the Royal
Society B: Biological Sciences 361, 1417–1432 (2006).
319. Roca, M., Moliner, V., Tuñón, I. & Hynes, J. T. Coupling between Protein and
Reaction Dynamics in Enzymatic Processes: Application of Grote−Hynes Theory to
Catechol O-Methyltransferase. J. Am. Chem. Soc. 128, 6186–6193 (2006).
320. Ruiz-Pernía, J. J., Tuñón, I., Moliner, V., Hynes, J. T. & Roca, M. Dynamic Effects
on Reaction Rates in a Michael Addition Catalyzed by Chalcone Isomerase. Beyond
the Frozen Environment Approach. J. Am. Chem. Soc. 130, 7477–7488 (2008).
321. Warshel, A. Dynamics of enzymatic reactions. PNAS 81, 444–448 (1984). 159
322. Simulations of quantum mechanical corrections for rate constants of hydride-
transfer reactions in enzymes and solutions | The Journal of Physical Chemistry.
https://pubs.acs.org/doi/abs/10.1021/j100175a009.
323. Eyring, H. The Activated Complex in Chemical Reactions. J. Chem. Phys. 3, 107–
115 (1935).
324. Warshel, A. Energetics of enzyme catalysis. PNAS 75, 5250–5254 (1978).
325. Warshel, A. Electrostatic Origin of the Catalytic Power of Enzymes and the Role of
Preorganized Active Sites. J. Biol. Chem. 273, 27035–27038 (1998).
326. Warshel, Arieh. & Russell, Stephen. Theoretical correlation of structure and
energetics in the catalytic reaction of trypsin. J. Am. Chem. Soc. 108, 6569–6579
(1986).
327. The mechanisms of hydrolysis of glycosides and their revelance to enzyme-
catalysed reactions | Proceedings of the Royal Society of London. Series B.
Biological Sciences.
https://royalsocietypublishing.org/doi/abs/10.1098/rspb.1967.0036?casa_token=NU
wG8rSCK-wAAAAA:1EbFsq-HzqnaCrqklR-
9FWk5hgXMw52bo78WxrscF2GQximbq36X84FGyAw4ObgrawwBpLRfViNuLn
M.
328. Hwang, J. K., Chu, Z. T., Yadav, A. & Warshel, A. Simulations of quantum
mechanical corrections for rate constants of hydride-transfer reactions in enzymes
and solutions. J. Phys. Chem. 95, 8445–8448 (1991).
329. Schramm, V. L. Transition States and Transition State Analogue Interactions with
Enzymes. Acc Chem Res 48, 1032–1039 (2015). 160
330. Pisliakov, A. V., Cao, J., Kamerlin, S. C. L. & Warshel, A. Enzyme millisecond
conformational dynamics do not catalyze the chemical step. PNAS 106, 17359–
17364 (2009).
331. Carrico, I. S. Chemoselective modification of proteins: hitting the target. Chem. Soc.
Rev. 37, 1423–1431 (2008).
332. Gamblin, D. P., van Kasteren, S. I., Chalker, J. M. & Davis, B. G. Chemical
approaches to mapping the function of post-translational modifications. FEBS J.
275, 1949–1959 (2008).
333. Walsh, C. Posttranslational Modification of Proteins: Expanding Nature’s
Inventory. (Roberts and Company Publishers, 2006).
334. Walsh, C. T., Garneau‐Tsodikova, S. & Gatto, G. J. Posttranslationale
Proteinmodifikation: die Chemie der Proteomdiversifizierung. Angewandte Chemie
117, 7508–7539 (2005).
335. Marino, S. M. & Gladyshev, V. N. Analysis and Functional Prediction of Reactive
Cysteine Residues. J. Biol. Chem. 287, 4419–4425 (2012).
336. Miseta, A. & Csutora, P. Relationship between the occurrence of cysteine in
proteins and the complexity of organisms. Mol. Biol. Evol. 17, 1232–1239 (2000).
337. Fomenko, D. E., Marino, S. M. & Gladyshev, V. N. Functional Diversity of
Cysteine Residues in Proteins and Unique Features of Catalytic Redox-active
Cysteines in Thiol Oxidoreductases. Mol Cells 26, 228–235 (2008).
338. Biswas, S., Chida, A. S. & Rahman, I. Redox modifications of protein-thiols:
emerging roles in cell signaling. Biochem. Pharmacol. 71, 551–564 (2006). 161
339. Gunnoo, S. B. & Madder, A. Chemical Protein Modification through Cysteine.
ChemBioChem 17, 529–553 (2016).
340. Foley, T. L. & Burkart, M. D. Site-specific protein modification: advances and
applications. Curr Opin Chem Biol 11, 12–19 (2007).
341. Bonham, C. A. & Vacratsis, P. O. Redox Regulation of the Human Dual Specificity
Phosphatase YVH1 through Disulfide Bond Formation. J Biol Chem 284, 22853–
22864 (2009).
342. Winterbourn, C. C. & Hampton, M. B. Thiol chemistry and specificity in redox
signaling. Free Radical Biology and Medicine 45, 549–561 (2008).
343. Devasagayam, T. P. A. et al. Free radicals and antioxidants in human health: current
status and future prospects. J Assoc Physicians India 52, 794–804 (2004).
344. Nagy, P. Kinetics and mechanisms of thiol-disulfide exchange covering direct
substitution and thiol oxidation-mediated pathways. Antioxid. Redox Signal. 18,
1623–1641 (2013).
345. Winther, J. R. & Thorpe, C. Quantification of thiols and disulfides. Biochimica et
Biophysica Acta (BBA) - General Subjects 1840, 838–846 (2014).
346. Poole, L. B. The Basics of Thiols and Cysteines in Redox Biology and Chemistry.
Free Radic Biol Med 0, 148–157 (2015).
347. Guttmann, R. P. & Powell, T. J. Redox Regulation of Cysteine-Dependent Enzymes
in Neurodegeneration. International Journal of Cell Biology
https://www.hindawi.com/journals/ijcb/2012/703164/ (2012)
doi:10.1155/2012/703164. 162
348. Reddie, K. G. & Carroll, K. S. Expanding the functional diversity of proteins
through cysteine oxidation. Current Opinion in Chemical Biology 12, 746–754
(2008).
349. Ahmad, S. et al. Protein oxidation: an overview of metabolism of sulphur
containing amino acid, cysteine. Front Biosci (Schol Ed) 9, 71–87 (2017).
350. Chung, H. S., Wang, S.-B., Venkatraman, V., Murray, C. I. & Van Eyk, J. E.
Cysteine Oxidative Post-translational Modifications: Emerging Regulation in the
Cardiovascular System. Circ Res 112, 382–392 (2013).
351. Zeida, A. et al. Thiol redox biochemistry: insights from computer simulations.
Biophys Rev 6, 27–46 (2014).
352. Fu, P. P., Xia, Q., Hwang, H.-M., Ray, P. C. & Yu, H. Mechanisms of nanotoxicity:
Generation of reactive oxygen species. Journal of Food and Drug Analysis 22, 64–
75 (2014).
353. Sarniak, A., Lipińska, J., Tytman, K. & Lipińska, S. Endogenous mechanisms of
reactive oxygen species (ROS) generation. Postepy Hig Med Dosw (Online) 70,
1150–1165 (2016).
354. Bigarella, C. L., Liang, R. & Ghaffari, S. Stem cells and the impact of ROS
signaling. Development 141, 4206–4218 (2014).
355. Newsholme, P., Cruzat, V. F., Keane, K. N., Carlessi, R. & de Bittencourt, P. I. H.
Molecular mechanisms of ROS production and oxidative stress in diabetes.
Biochem. J. 473, 4527–4550 (2016).
356. Dalle-Donne, I. et al. Molecular mechanisms and potential clinical significance of
S-glutathionylation. Antioxid. Redox Signal. 10, 445–473 (2008). 163
357. Moran, L. K., Gutteridge, J. M. & Quinlan, G. J. Thiols in cellular redox signalling
and control. Curr. Med. Chem. 8, 763–772 (2001).
358. Taylor, E. R. et al. Reversible glutathionylation of complex I increases
mitochondrial superoxide formation. J. Biol. Chem. 278, 19603–19610 (2003).
359. Bindoli, A., Fukuto, J. M. & Forman, H. J. Thiol Chemistry in Peroxidase Catalysis
and Redox Signaling. Antioxid Redox Signal 10, 1549–1564 (2008).
360. Lin, H.-H. & Tseng, L.-Y. DBCP: a web server for disulfide bonding connectivity
pattern prediction without the prior knowledge of the bonding state of cysteines.
Nucleic Acids Res 38, W503–W507 (2010).
361. Paget, M. S. B. & Buttner, M. J. Thiol-Based Regulatory Switches. Annu. Rev.
Genet. 37, 91–121 (2003).
362. Chung Heaseung S., Wang Sheng-Bing, Venkatraman Vidya, Murray Christopher I.
& Van Eyk Jennifer E. Cysteine Oxidative Posttranslational Modifications.
Circulation Research 112, 382–392 (2013).
363. He, H. T. et al. Synthesis and chemical stability of a disulfide bond in a model
cyclic pentapeptide: Cyclo(1,4)‐Cys‐Gly‐Phe‐Cys‐Gly‐OH. Journal of
Pharmaceutical Sciences 95, 2222–2234 (2006).
364. Bulleid, N. J. Disulfide Bond Formation in the Mammalian Endoplasmic Reticulum.
Cold Spring Harb Perspect Biol 4, (2012).
365. Gilbert, H. F. [2] Thiol/disulfide exchange equilibria and disulfidebond stability. in
Methods in Enzymology vol. 251 8–28 (Academic Press, 1995). 164
366. THING, M., ZHANG, J., LAURENCE, J. & TOPP, E. M. Thiol-Disulfide
Interchange in the Tocinoic Acid/Glutathione System During Freezing and Drying.
J Pharm Sci 99, 4849–4856 (2010).
367. Kolšek, K., Aponte-Santamaría, C. & Gräter, F. Accessibility explains preferred
thiol-disulfide isomerization in a protein domain. Scientific Reports 7, 1–10 (2017).
368. Schwertassek, U. et al. Selective redox regulation of cytokine receptor signaling by
extracellular thioredoxin-1. EMBO J. 26, 3086–3097 (2007).
369. Singh, R. & Whitesides, G. M. Thiol-disulfide interchange. in Sulphur-Containing
Functional Groups (1993) (eds. Patai, S. & Rappoport, Z.) 633–658 (John Wiley &
Sons, Inc., 1993). doi:10.1002/9780470034408.ch13.
370. Riemer, J., Bulleid, N. & Herrmann, J. M. Disulfide Formation in the ER and
Mitochondria: Two Solutions to a Common Process. Science 324, 1284–1287
(2009).
371. Bach, R. D., Dmitrenko, O. & Thorpe, C. Mechanism of Thiolate−Disulfide
Interchange Reactions in Biochemistry. J. Org. Chem. 73, 12–21 (2008).
372. Depuydt, M., Messens, J. & Collet, J.-F. How proteins form disulfide bonds.
Antioxid. Redox Signal. 15, 49–66 (2011).
373. Bulleid, N. J. & Ellgaard, L. Multiple ways to make disulfides. Trends in
Biochemical Sciences 36, 485–492 (2011).
374. Messens, J. & Collet, J.-F. Pathways of disulfide bond formation in Escherichia coli.
Int. J. Biochem. Cell Biol. 38, 1050–1062 (2006). 165
375. Ziegler, D. M. Role of Reversible Oxidation-Reduction of Enzyme Thiols-
Disulfides in Metabolic Regulation. Annual Review of Biochemistry 54, 305–329
(1985).
376. Zheng, M., Åslund, F. & Storz, G. Activation of the OxyR Transcription Factor by
Reversible Disulfide Bond Formation. Science 279, 1718–1722 (1998).
377. Landeta, C., Boyd, D. & Beckwith, J. Disulfide bond formation in prokaryotes.
Nature Microbiology 3, 270–280 (2018).
378. Dröge, W. Aging-related changes in the thiol/disulfide redox state: implications for
the use of thiol antioxidants. Experimental Gerontology 37, 1333–1345 (2002).
379. Go Young-Mi & Jones Dean P. Intracellular Proatherogenic Events and Cell
Adhesion Modulated by Extracellular Thiol/Disulfide Redox State. Circulation 111,
2973–2980 (2005).
380. Dickinson, D. A. & Forman, H. J. Cellular glutathione and thiols metabolism.
Biochemical Pharmacology 64, 1019–1026 (2002).
381. Moran, L. K. & Quinlan, J. M. C. G. and G. J. Thiols in Cellular Redox Signalling
and Control. Current Medicinal Chemistry
http://www.eurekaselect.com/65361/article (2001).
382. Go, Y.-M. & Jones, D. P. Thiol/disulfide redox states in signaling and sensing. Crit.
Rev. Biochem. Mol. Biol. 48, 173–181 (2013).
383. Cysteine/cystine couple is a newly recognized node in the circuitry for biologic
redox signaling and control | The FASEB Journal.
https://www.fasebj.org/doi/abs/10.1096/fj.03-0971fje. 166
384. Jonas, C. R., Ziegler, T. R., Gu, L. i H. & Jones, D. P. Extracellular thiol/disulfide
redox state affects proliferation rate in a human colon carcinoma (Caco2) cell line.
Free Radical Biology and Medicine 33, 1499–1506 (2002).
385. Jorgenson, T. C., Zhong, W. & Oberley, T. D. Redox Imbalance and Biochemical
Changes in Cancer. Cancer Res 73, 6118–6123 (2013).
386. Giles, G. I. The Redox Regulation of Thiol Dependent Signaling Pathways in
Cancer.
https://www.ingentaconnect.com/content/ben/cpd/2006/00000012/00000034/art000
03 (2006) doi:info:doi/10.2174/138161206779010549.
387. Shan, W., Zhong, W., Zhao, R. & Oberley, T. D. Thioredoxin 1 as a subcellular
biomarker of redox imbalance in human prostate cancer progression. Free Radical
Biology and Medicine 49, 2078–2087 (2010).
388. Chen, V. M. et al. Evidence for Activation of Tissue Factor by an Allosteric
Disulfide Bond. Biochemistry 45, 12020–12028 (2006).
389. Hart, E. J. & Powers-Lee, S. G. Role of Cys-1327 and Cys-1337 in Redox
Sensitivity and Allosteric Monitoring in Human Carbamoyl Phosphate Synthetase.
J. Biol. Chem. 284, 5977–5985 (2009).
390. Crystal Structures of the Reduced, Sulfenic Acid, and Mixed Disulfide Forms of
SarZ, a Redox Active Global Regulator in Staphylococcus aureus.
http://www.jbc.org/content/284/35/23517.
391. Cook, K. M., McNeil, H. P. & Hogg, P. J. Allosteric Control of βII-Tryptase by a
Redox Active Disulfide Bond. J. Biol. Chem. 288, 34920–34929 (2013). 167
392. Rajpal, G. & Arvan, P. Chapter 236 - Disulfide Bond Formation. in Handbook of
Biologically Active Peptides (Second Edition) (ed. Kastin, A. J.) 1721–1729
(Academic Press, 2013). doi:10.1016/B978-0-12-385095-9.00236-0.
393. Hatahet, F., Boyd, D. & Beckwith, J. Disulfide Bond Formation in Prokaryotes:
History, Diversity and Design. Biochim Biophys Acta 1844, 1402–1414 (2014).
394. Chen, W., Helenius, J., Braakman, I. & Helenius, A. Cotranslational folding and
calnexin binding during glycoprotein synthesis. Proc Natl Acad Sci U S A 92, 6229–
6233 (1995).
395. Ushioda, R. et al. ERdj5 is required as a disulfide reductase for degradation of
misfolded proteins in the ER. Science 321, 569–572 (2008).
396. Darby, N. J., Freedman, R. B. & Creighton, T. E. Dissecting the mechanism of
protein disulfide isomerase: catalysis of disulfide bond formation in a model
peptide. Biochemistry 33, 7937–7947 (1994).
397. Laboissière, M. C. A., Sturley, S. L. & Raines, R. T. The Essential Function of
Protein-disulfide Isomerase Is to Unscramble Non-native Disulfide Bonds. J. Biol.
Chem. 270, 28006–28009 (1995).
398. Appenzeller-Herzog, C. & Ellgaard, L. In Vivo Reduction-Oxidation State of
Protein Disulfide Isomerase: The Two Active Sites Independently Occur in the
Reduced and Oxidized Forms. Antioxidants & Redox Signaling 10, 55–64 (2007).
399. Bardwell, J. C. et al. A pathway for disulfide bond formation in vivo. PNAS 90,
1038–1042 (1993).
400. Ito, K. & Inaba, K. The disulfide bond formation (Dsb) system. Current Opinion in
Structural Biology 18, 450–458 (2008). 168
401. Holmgren, A. Enzymatic reduction-oxidation of protein disulfides by thioredoxin. in
Methods in Enzymology vol. 107 295–300 (Academic Press, 1984).
402. Hillson, D. A., Lambert, N. & Freedman, R. B. Formation and isomerization of
disulfide bonds in proteins: Protein disulfide-isomerase. in Methods in Enzymology
vol. 107 281–294 (Academic Press, 1984).
403. Bardwell, J. C. A., McGovern, K. & Beckwith, J. Identification of a protein required
for disulfide bond formation in vivo. Cell 67, 581–589 (1991).
404. Riemer, J. et al. A luminal flavoprotein in endoplasmic reticulum-associated
degradation. Proc. Natl. Acad. Sci. U.S.A. 106, 14831–14836 (2009).
405. Kadokura, H., Katzen, F. & Beckwith, J. Protein disulfide bond formation in
prokaryotes. Annu. Rev. Biochem. 72, 111–135 (2003).
406. Cho, S.-H., Porat, A., Ye, J. & Beckwith, J. Redox-active cysteines of a membrane
electron transporter DsbD show dual compartment accessibility. EMBO J. 26,
3509–3520 (2007).
407. Segatori, L., Paukstelis, P. J., Gilbert, H. F. & Georgiou, G. Engineered DsbC
chimeras catalyze both protein oxidation and disulfide-bond isomerization in
Escherichia coli: Reconciling two competing pathways. Proc. Natl. Acad. Sci.
U.S.A. 101, 10018–10023 (2004).
408. Acids (Sulfonic, Sulfinic, Sulfenic) and Derivatives. in Nomenclature of Organic
Compounds vol. 126 146–152 (AMERICAN CHEMICAL SOCIETY, 1974).
409. van Bergen, L. A. H., Roos, G. & De Proft, F. From thiol to sulfonic acid: modeling
the oxidation pathway of protein thiols by hydrogen peroxide. J Phys Chem A 118,
6078–6084 (2014). 169
410. Lo Conte, M. & Carroll, K. S. The Redox Biochemistry of Protein Sulfenylation and
Sulfinylation. J. Biol. Chem. 288, 26480–26488 (2013).
411. Poole, L. B., Karplus, P. A. & Claiborne, A. Protein sulfenic acids in redox
signaling. Annu. Rev. Pharmacol. Toxicol. 44, 325–347 (2004).
412. Carballal, S. et al. Sulfenic acid in human serum albumin. Amino Acids 32, 543–551
(2007).
413. Heppner, D. E., Janssen-Heininger, Y. M. W. & van der Vliet, A. The role of
sulfenic acids in cellular redox signaling: Reconciling chemical kinetics and
molecular detection strategies. Arch. Biochem. Biophys. 616, 40–46 (2017).
414. Saurin, A. T., Neubert, H., Brennan, J. P. & Eaton, P. Widespread sulfenic acid
formation in tissues in response to hydrogen peroxide. PNAS 101, 17982–17987
(2004).
415. Simmons, C. R. et al. Crystal Structure of Mammalian Cysteine Dioxygenase A
NOVEL MONONUCLEAR IRON CENTER FOR CYSTEINE THIOL
OXIDATION. J. Biol. Chem. 281, 18723–18733 (2006).
416. Formation and reactions of sulfenic acids in proteins | Accounts of Chemical
Research. https://pubs.acs.org/doi/abs/10.1021/ar50104a003.
417. Rehder, D. S. & Borges, C. R. Cysteine sulfenic Acid as an Intermediate in
Disulfide Bond Formation and Nonenzymatic Protein Folding. Biochemistry 49,
7748–7755 (2010).
418. Lee, J.-W., Soonsanga, S. & Helmann, J. D. A complex thiolate switch regulates the
Bacillus subtilis organic peroxide sensor OhrR. PNAS 104, 8743–8748 (2007). 170
419. Redox regulation of protein tyrosine phosphatase 1B involves a sulphenyl-amide
intermediate | Nature. https://www.nature.com/articles/nature01680.
420. Salmeen, A. et al. Redox regulation of protein tyrosine phosphatase 1B involves a
sulphenyl-amide intermediate. Nature 423, 769–773 (2003).
421. van Montfort, R. L. M., Congreve, M., Tisi, D., Carr, R. & Jhoti, H. Oxidation state
of the active-site cysteine in protein tyrosine phosphatase 1B. Nature 423, 773–777
(2003).
422. Depuydt, M. et al. A Periplasmic Reducing System Protects Single Cysteine
Residues from Oxidation. Science 326, 1109–1111 (2009).
423. Torchinskii, Y. M. Sulfhydryl and Disulfide Groups of Proteins. (Springer Science
& Business Media, 2013).
424. Kice, J. L. Mechanisms and Reactivity in Reactions of Organic Oxyacids of Sulfur
and their Anhydrides. in Advances in Physical Organic Chemistry (eds. Gold, V. &
Bethell, D.) vol. 17 65–181 (Academic Press, 1980).
425. Claiborne, A., Miller, H., Parsonage, D. & Ross, R. P. Protein-sulfenic acid
stabilization and function in enzyme catalysis and gene regulation. FASEB J. 7,
1483–1490 (1993).
426. Klomsiri, C., Karplus, P. A. & Poole, L. B. Cysteine-Based Redox Switches in
Enzymes. Antioxid Redox Signal 14, 1065–1077 (2011).
427. Kaplan, N. et al. Localized cysteine sulfenic acid formation by vascular endothelial
growth factor: role in endothelial cell migration and angiogenesis. Free Radic. Res.
45, 1124–1135 (2011). 171
428. Peroxide-dependent sulfenylation of the EGFR catalytic site enhances kinase
activity | Nature Chemical Biology. https://www.nature.com/articles/nchembio.736.
429. Paulsen, C. E. & Carroll, K. S. Orchestrating Redox Signaling Networks through
Regulatory Cysteine Switches. ACS Chem. Biol. 5, 47–62 (2010).
430. Xu, D., Rovira, I. I. & Finkel, T. Oxidants Painting the Cysteine Chapel: Redox
Regulation of PTPs. Developmental Cell 2, 251–252 (2002).
431. Peshenko, I. V. & Shichi, H. Oxidation of active center cysteine of bovine 1-Cys
peroxiredoxin to the cysteine sulfenic acid form by peroxide and peroxynitrite. Free
Radical Biology and Medicine 31, 292–303 (2001).
432. Turell, L., Carballal, S., Botti, H., Radi, R. & Alvarez, B. Oxidation of the albumin
thiol to sulfenic acid and its implications in the intravascular compartment.
Brazilian Journal of Medical and Biological Research 42, 305–311 (2009).
433. Turell, L. et al. Reactivity of Sulfenic Acid in Human Serum Albumin.
Biochemistry 47, 358–367 (2008).
434. Michalek, R. D. et al. The Requirement of Reversible Cysteine Sulfenic Acid
Formation for T Cell Activation and Function. The Journal of Immunology 179,
6456–6467 (2007).
435. Roos, G. & Messens, J. Protein sulfenic acid formation: From cellular damage to
redox regulation. Free Radical Biology and Medicine 51, 314–326 (2011).
436. Rabilloud, T. et al. Proteomics analysis of cellular response to oxidative stress.
Evidence for in vivo overoxidation of peroxiredoxins at their active site. J. Biol.
Chem. 277, 19396–19401 (2002). 172
437. Wood, Z. A., Poole, L. B. & Karplus, P. A. Peroxiredoxin Evolution and the
Regulation of Hydrogen Peroxide Signaling. Science 300, 650–653 (2003).
438. Yang, K.-S. et al. Inactivation of human peroxiredoxin I during catalysis as the
result of the oxidation of the catalytic site cysteine to cysteine-sulfinic acid. J. Biol.
Chem. 277, 38029–38036 (2002).
439. The Parkinson’s disease protein DJ-1 is neuroprotective due to cysteine-sulfinic
acid-driven mitochondrial localization | PNAS.
https://www.pnas.org/content/101/24/9103.short.
440. Cysteine pKa Depression by a Protonated Glutamic Acid in Human DJ-1 |
Biochemistry. https://pubs.acs.org/doi/abs/10.1021/bi800282d.
441. Formation of a Stabilized Cysteine Sulfinic Acid Is Critical for the Mitochondrial
Function of the Parkinsonism Protein DJ-1.
http://www.jbc.org/content/284/10/6476.short.
442. Wilson, M. A. The Role of Cysteine Oxidation in DJ-1 Function and Dysfunction.
Antioxidants & Redox Signaling 15, 111–122 (2010).
443. Miyanaga, A., Fushinobu, S., Ito, K. & Wakagi, T. Crystal Structure of Cobalt-
Containing Nitrile Hydratase. Biochemical and Biophysical Research
Communications 288, 1169–1174 (2001).
444. Arakawa, T. et al. Structural Basis for Catalytic Activation of Thiocyanate
Hydrolase Involving Metal-Ligated Cysteine Modification. J. Am. Chem. Soc. 131,
14838–14843 (2009).
445. Biteau, B., Labarre, J. & Toledano, M. B. ATP-dependent reduction of cysteine-
sulphinic acid by S. cerevisiae sulphiredoxin. Nature 425, 980–984 (2003). 173
446. Woo, H. A. et al. Reversing the inactivation of peroxiredoxins caused by cysteine
sulfinic acid formation. Science 300, 653–656 (2003).
447. Melino, G. et al. S -nitrosylation regulates apoptosis. Nature 388, 432–433 (1997).
448. Woo, H. A. et al. Reduction of Cysteine Sulfinic Acid by Sulfiredoxin Is Specific to
2-Cys Peroxiredoxins. J. Biol. Chem. 280, 3125–3128 (2005).
449. Chang, T.-S. et al. Characterization of Mammalian Sulfiredoxin and Its Reactivation
of Hyperoxidized Peroxiredoxin through Reduction of Cysteine Sulfinic Acid in the
Active Site to Cysteine. J. Biol. Chem. 279, 50994–51001 (2004).
450. Jeong, W., Park, S. J., Chang, T.-S., Lee, D.-Y. & Rhee, S. G. Molecular
Mechanism of the Reduction of Cysteine Sulfinic Acid of Peroxiredoxin to Cysteine
by Mammalian Sulfiredoxin. J. Biol. Chem. 281, 14400–14407 (2006).
451. Stressin’ Sestrins take an aging fight | EMBO Molecular Medicine.
https://www.embopress.org/doi/full/10.1002/emmm.201000097.
452. Budanov, A. V., Sablina, A. A., Feinstein, E., Koonin, E. V. & Chumakov, P. M.
Regeneration of peroxiredoxins by p53-regulated sestrins, homologs of bacterial
AhpD. Science 304, 596–600 (2004).
453. Lim, J. C. et al. Irreversible Oxidation of the Active-site Cysteine of Peroxiredoxin
to Cysteine Sulfonic Acid for Enhanced Molecular Chaperone Activity. J. Biol.
Chem. 283, 28873–28880 (2008).
454. Wagner, E. et al. A method for detection of overoxidation of cysteines:
peroxiredoxins are oxidized in vivo at the active-site cysteine during oxidative
stress. Biochem J 366, 777–785 (2002). 174
455. Miki, H. & Funato, Y. Regulation of intracellular signalling through cysteine
oxidation by reactive oxygen species. J Biochem 151, 255–261 (2012).
456. Östman, A., Frijhoff, J., Sandin, Å. & Böhmer, F.-D. Regulation of protein tyrosine
phosphatases by reversible oxidation. J Biochem 150, 345–356 (2011).
457. Tanner, J. J., Parsons, Z. D., Cummings, A. H., Zhou, H. & Gates, K. S. Redox
Regulation of Protein Tyrosine Phosphatases: Structural and Chemical Aspects.
Antioxidants & Redox Signaling 15, 77–97 (2010).
458. von, M. C. et al. Singlet oxygen inactivates protein tyrosine phosphatase-1B by
oxidation of the active site cysteine. Biological Chemistry 387, 1399–1404 (2006).
459. Lee, S.-R., Kwon, K.-S., Kim, S.-R. & Rhee, S. G. Reversible Inactivation of
Protein-tyrosine Phosphatase 1B in A431 Cells Stimulated with Epidermal Growth
Factor. J. Biol. Chem. 273, 15366–15372 (1998).
460. Kemble, D. J. & Sun, G. Direct and specific inactivation of protein tyrosine kinases
in the Src and FGFR families by reversible cysteine oxidation. PNAS 106, 5070–
5075 (2009).
461. Truong, T. H. & Carroll, K. S. Redox Regulation of Epidermal Growth Factor
Receptor Signaling through Cysteine Oxidation. Biochemistry 51, 9954–9965
(2012).
462. Holmgren, A. Thioredoxin structure and mechanism: conformational changes on
oxidation of the active-site sulfhydryls to a disulfide. Structure 3, 239–243 (1995).
463. Prongay, A. J. & Williams, C. H. Oxidation-reduction properties of Escherichia coli
thioredoxin reductase altered at each active site cysteine residue. J. Biol. Chem. 267,
25181–25188 (1992). 175
464. Saitoh, M. et al. Mammalian thioredoxin is a direct inhibitor of apoptosis signal-
regulating kinase (ASK) 1. The EMBO Journal 17, 2596–2606 (1998).
465. Ichijo, H. et al. Induction of Apoptosis by ASK1, a Mammalian MAPKKK That
Activates SAPK/JNK and p38 Signaling Pathways. Science 275, 90–94 (1997).
466. Caliceti, C., Nigro, P., Rizzo, P. & Ferrari, R. ROS, Notch, and Wnt Signaling
Pathways: Crosstalk between Three Major Regulators of Cardiovascular Biology.
Biomed Res Int 2014, (2014).
467. Funato, Y. & Miki, H. Nucleoredoxin, a Novel Thioredoxin Family Member
Involved in Cell Growth and Differentiation. Antioxidants & Redox Signaling 9,
1035–1058 (2007).
468. Funato, Y. et al. Nucleoredoxin Sustains Wnt/β-Catenin Signaling by Retaining a
Pool of Inactive Dishevelled Protein. Current Biology 20, 1945–1952 (2010).
469. Kurooka, H. et al. Cloning and Characterization of the Nucleoredoxin Gene That
Encodes a Novel Nuclear Protein Related to Thioredoxin. Genomics 39, 331–339
(1997).
470. Steitz, S. A., Tsang, M. & Sussman, D. J. WNT-mediated relocalization of
dishevelled proteins. In Vitro Cell.Dev.Biol.-Animal 32, 441–445 (1996).
471. Woo, H. A. et al. Reversible Oxidation of the Active Site Cysteine of
Peroxiredoxins to Cysteine Sulfinic Acid IMMUNOBLOT DETECTION WITH
ANTIBODIES SPECIFIC FOR THE HYPEROXIDIZED CYSTEINE-
CONTAINING SEQUENCE. J. Biol. Chem. 278, 47361–47364 (2003).
472. Hall, A., Karplus, P. A. & Poole, L. B. Typical 2-Cys Peroxiredoxins: Structures,
mechanisms and functions. FEBS J 276, 2469–2477 (2009). 176
473. Rhee, S. G. Overview on Peroxiredoxin. Mol Cells 39, 1–5 (2016).
474. Brandstaedter, C. et al. The interactome of 2-Cys peroxiredoxins in Plasmodium
falciparum. Scientific Reports 9, 1–15 (2019).
475. Netto, L. E. S. & Antunes, F. The Roles of Peroxiredoxin and Thioredoxin in
Hydrogen Peroxide Sensing and in Signal Transduction. Mol Cells 39, 65–71
(2016).
476. Rhee, S. G., Woo, H. A., Kil, I. S. & Bae, S. H. Peroxiredoxin Functions as a
Peroxidase and a Regulator and Sensor of Local Peroxides. J. Biol. Chem. 287,
4403–4410 (2012).
477. Jang, H. H. et al. Two Enzymes in One: Two Yeast Peroxiredoxins Display
Oxidative Stress-Dependent Switching from a Peroxidase to a Molecular Chaperone
Function. Cell 117, 625–635 (2004).
478. Oligomeric peroxiredoxin-I is an essential intermediate for p53 to activate MST1
kinase and apoptosis | Oncogene. https://www.nature.com/articles/onc2011139.
479. Rhee, S. G., Chae, H. Z. & Kim, K. Peroxiredoxins: A historical overview and
speculative preview of novel mechanisms and emerging concepts in cell signaling.
Free Radical Biology and Medicine 38, 1543–1552 (2005).
480. Leichert, L. I. et al. Quantifying changes in the thiol redox proteome upon oxidative
stress in vivo. PNAS 105, 8197–8202 (2008).
481. Colussi, C. et al. H2O2-induced block of glycolysis as an active ADP-ribosylation
reaction protecting cells from apoptosis. The FASEB Journal 14, 2266–2276 (2000). 177
482. Hess, D. T., Matsumoto, A., Kim, S.-O., Marshall, H. E. & Stamler, J. S. Protein S -
nitrosylation: purview and parameters. Nature Reviews Molecular Cell Biology 6,
150–166 (2005).
483. xloa004.PDF | Elsevier Enhanced Reader.
https://reader.elsevier.com/reader/sd/pii/S0003986185712313?token=550C3667065
8CCA6AE7A55D5E28FAFF4FCF55A0CF9B42807FA80D8AC8F34F6577F11C6
1675E1BCC2A9541B7B3A39080C doi:10.1006/abbi.1995.1231.
484. Hess, D. T., Matsumoto, A., Nudelman, R. & Stamler, J. S. S -nitrosylation:
spectrum and specificity. Nature Cell Biology 3, E46–E48 (2001).
485. Lima Brian, Forrester Michael T., Hess Douglas T. & Stamler Jonathan S. S-
Nitrosylation in Cardiovascular Signaling. Circulation Research 106, 633–646
(2010).
486. Foster, M. W., McMahon, T. J. & Stamler, J. S. S-nitrosylation in health and
disease. Trends in Molecular Medicine 9, 160–168 (2003).
487. Chen, Y.-Y. et al. Cysteine S-Nitrosylation Protects Protein-tyrosine Phosphatase
1B against Oxidation-induced Permanent Inactivation. J. Biol. Chem. 283, 35265–
35272 (2008).
488. Ridnour, L. A. et al. The chemistry of nitrosative stress induced by nitric oxide and
reactive nitrogen oxide species. Putting perspective on stressful biological
situations. Biol. Chem. 385, 1–10 (2004).
489. Broillet, M.-C. S-Nitrosylation of proteins. CMLS, Cell. Mol. Life Sci. 55, 1036
(1999). 178
490. Martínez-Ruiz, A. & Lamas, S. S-nitrosylation: a potential new paradigm in signal
transduction. Cardiovasc. Res. 62, 43–52 (2004).
491. Puyaubert, J., Fares, A., Rézé, N., Peltier, J.-B. & Baudouin, E. Identification of
endogenously S-nitrosylated proteins in Arabidopsis plantlets: Effect of cold stress
on cysteine nitrosylation level. Plant Science 215–216, 150–156 (2014).
492. Persichini, T., Colasanti, M., Lauro, G. M. & Ascenzi, P. Cysteine Nitrosylation
Inactivates the HIV-1 Protease. Biochemical and Biophysical Research
Communications 250, 575–576 (1998).
493. Yoshida, T. et al. Nitric oxide activates TRP channels by cysteine S-nitrosylation.
Nat Chem Biol 2, 596–607 (2006).
494. Stamler, J. S., Lamas, S. & Fang, F. C. Nitrosylation: The Prototypic Redox-Based
Signaling Mechanism. Cell 106, 675–683 (2001).
495. Shelton, M. D., Chock, P. B. & Mieyal, J. J. Glutaredoxin: Role in Reversible
Protein S-Glutathionylation and Regulation of Redox Signal Transduction and
Protein Translocation. Antioxidants & Redox Signaling 7, 348–366 (2005).
496. Grek, C. L., Zhang, J., Manevich, Y., Townsend, D. M. & Tew, K. D. Causes and
Consequences of Cysteine S-Glutathionylation. J. Biol. Chem. 288, 26497–26504
(2013).
497. Xiong, Y., Uys, J. D., Tew, K. D. & Townsend, D. M. S-Glutathionylation: From
Molecular Mechanisms to Health Outcomes. Antioxid Redox Signal 15, 233–270
(2011). 179
498. Adachi, T. et al. S-glutathiolation of Ras mediates redox-sensitive signaling by
angiotensin II in vascular smooth muscle cells. J. Biol. Chem. 279, 29857–29862
(2004).
499. Townsend, D. M. et al. A glutathione S-transferase pi-activated prodrug causes
kinase activation concurrent with S-glutathionylation of proteins. Mol. Pharmacol.
69, 501–508 (2006).
500. Barrett, W. C. et al. Regulation of PTP1B via glutathionylation of the active site
cysteine 215. Biochemistry 38, 6699–6705 (1999).
501. Humphries, K. M., Juliano, C. & Taylor, S. S. Regulation of cAMP-dependent
protein kinase activity by glutathionylation. J. Biol. Chem. 277, 43505–43511
(2002).
502. Yu, C.-X., Li, S. & Whorton, A. R. Redox regulation of PTEN by S-nitrosothiols.
Mol. Pharmacol. 68, 847–854 (2005).
503. Cruz, C. M. et al. ATP activates a reactive oxygen species-dependent oxidative
stress response and secretion of proinflammatory cytokines in macrophages. J. Biol.
Chem. 282, 2871–2879 (2007).
504. Rao, R. K. & Clayton, L. W. Regulation of protein phosphatase 2A by hydrogen
peroxide and glutathionylation. Biochem. Biophys. Res. Commun. 293, 610–616
(2002).
505. Velu, C. S., Niture, S. K., Doneanu, C. E., Pattabiraman, N. & Srivenugopal, K. S.
Human p53 is inhibited by glutathionylation of cysteines present in the proximal
DNA-binding domain during oxidative stress. Biochemistry 46, 7765–7780 (2007). 180
506. Karin, M. & Lin, A. NF-kappaB at the crossroads of life and death. Nat. Immunol.
3, 221–227 (2002).
507. Reynaert, N. L. et al. Dynamic redox control of NF-kappaB through glutaredoxin-
regulated S-glutathionylation of inhibitory kappaB kinase beta. Proc. Natl. Acad.
Sci. U.S.A. 103, 13086–13091 (2006).
508. Qanungo, S., Starke, D. W., Pai, H. V., Mieyal, J. J. & Nieminen, A.-L. Glutathione
supplementation potentiates hypoxic apoptosis by S-glutathionylation of p65-
NFkappaB. J. Biol. Chem. 282, 18427–18436 (2007).
509. Klatt, P., Molina, E. P. & Lamas, S. Nitric oxide inhibits c-Jun DNA binding by
specifically targeted S-glutathionylation. J. Biol. Chem. 274, 15857–15864 (1999).
510. Cross, J. V. & Templeton, D. J. Oxidative stress inhibits MEKK1 by site-specific
glutathionylation in the ATP-binding domain. Biochem. J. 381, 675–683 (2004).
511. Adler, V. et al. Regulation of JNK signaling by GSTp. EMBO J. 18, 1321–1334
(1999).
512. Townsend, D. M. et al. Novel role for glutathione S-transferase pi. Regulator of
protein S-Glutathionylation following oxidative and nitrosative stress. J. Biol.
Chem. 284, 436–445 (2009).
513. Wu, Y. et al. Human glutathione S-transferase P1-1 interacts with TRAF2 and
regulates TRAF2-ASK1 signals. Oncogene 25, 5787–5800 (2006).
514. Tew, K. D. Glutathione-associated enzymes in anticancer drug resistance. Cancer
Res. 54, 4313–4320 (1994). 181
515. Yin, Z., Ivanov, V. N., Habelhah, H., Tew, K. & Ronai, Z. Glutathione S-transferase
p elicits protection against H2O2-induced cell death via coordinated regulation of
stress kinases. Cancer Res. 60, 4053–4057 (2000).
516. Manevich, Y., Feinstein, S. I. & Fisher, A. B. Activation of the antioxidant enzyme
1-CYS peroxiredoxin requires glutathionylation mediated by heterodimerization
with pi GST. Proc. Natl. Acad. Sci. U.S.A. 101, 3780–3785 (2004).
517. Kim, Y.-J. et al. Prx1 suppresses radiation-induced c-Jun NH2-terminal kinase
signaling in lung cancer cells through interaction with the glutathione S-transferase
Pi/c-Jun NH2-terminal kinase complex. Cancer Res. 66, 7136–7142 (2006).
518. Mailloux, R. J., Jin, X. & Willmore, W. G. Redox regulation of mitochondrial
function with emphasis on cysteine oxidation reactions. Redox Biology 2, 123–139
(2014).
519. Chalker, J. M., Bernardes, G. J. L., Lin, Y. A. & Davis, B. G. Chemical
Modification of Proteins at Cysteine: Opportunities in Chemistry and Biology.
Chemistry – An Asian Journal 4, 630–640 (2009).
520. How Enzymes Work | Science.
https://science.sciencemag.org/content/320/5882/1428?casa_token=Az3CORVVbk
MAAAAA:iEvGg_uw1hrBlzoFmamSJf5qvZuYweI8TDphS5yuh1kqVDrGkVW2y
T17AkqJ_It0DIhhDjPnm5sRvZc.
521. Vögeli, B. & Erb, T. J. ‘Negative’ and ‘positive catalysis’: complementary
principles that shape the catalytic landscape of enzymes. Current Opinion in
Chemical Biology 47, 94–100 (2018). 182
522. Engineered control of enzyme structural dynamics and function - Boehr - 2018 -
Protein Science - Wiley Online Library.
https://onlinelibrary.wiley.com/doi/full/10.1002/pro.3379.
523. Henzler-Wildman, K. A. et al. Intrinsic motions along an enzymatic reaction
trajectory. Nature 450, 838–844 (2007).
524. Aviram, H. Y. et al. Direct observation of ultrafast large-scale dynamics of an
enzyme under turnover conditions. PNAS 115, 3243–3248 (2018).
525. Kim, T. H. et al. The role of dimer asymmetry and protomer dynamics in enzyme
catalysis. Science 355, eaag2355 (2017).
526. Fried, S. D. & Boxer, S. G. Electric Fields and Enzyme Catalysis. Annual Review of
Biochemistry 86, 387–415 (2017).
527. Warshel, A. & Bora, R. P. Perspective: Defining and quantifying the role of
dynamics in enzyme catalysis. J. Chem. Phys. 144, 180901 (2016).
528. Glowacki, D. R., Harvey, J. N. & Mulholland, A. J. Taking Ockham’s razor to
enzyme dynamics and catalysis. Nature Chem 4, 169–176 (2012).
529. Hanoian, P., Liu, C. T., Hammes-Schiffer, S. & Benkovic, S. Perspectives on
Electrostatics and Conformational Motions in Enzyme Catalysis. Acc. Chem. Res.
48, 482–489 (2015).
530. Conformational selection in protein binding and function - Weikl - 2014 - Protein
Science - Wiley Online Library.
https://onlinelibrary.wiley.com/doi/full/10.1002/pro.2539.
531. van den Bedem, H. & Fraser, J. S. Integrative, dynamic structural biology at atomic
resolution—it’s about time. Nature Methods 12, 307–318 (2015). 183
532. Danyal, K., Mayweather, D., Dean, D. R., Seefeldt, L. C. & Hoffman, B. M.
Conformational Gating of Electron Transfer from the Nitrogenase Fe Protein to
MoFe Protein. J. Am. Chem. Soc. 132, 6894–6895 (2010).
533. Kale, S. et al. Efficient coupling of catalysis and dynamics in the E1 component of
Escherichia coli pyruvate dehydrogenase multienzyme complex. Proceedings of the
National Academy of Sciences 105, 1158–1163 (2008).
534. Mehrabi, P. et al. Time-resolved crystallography reveals allosteric communication
aligned with molecular breathing. Science 365, 1167–1170 (2019).
535. Stiers, K. M. et al. Structural and dynamical description of the enzymatic reaction of
a phosphohexomutase. Structural Dynamics 6, 024703 (2019).
536. Pace, N. J. & Weerapana, E. Diverse Functional Roles of Reactive Cysteines. ACS
Chem. Biol. 8, 283–296 (2013).
537. Lonsdale, R. & Ward, R. A. Structure-based design of targeted covalent inhibitors.
Chem Soc Rev 47, 3816–3830 (2018).
538. Furnham, N., Blundell, T. L., DePristo, M. A. & Terwilliger, T. C. Is one solution
good enough? Nat. Struct. Mol. Biol. 13, 184–185; discussion 185 (2006).
539. Fraser, J. S. et al. Accessing protein conformational ensembles using room-
temperature X-ray crystallography. PNAS 108, 16247–16252 (2011).
540. Smith, J. L., Hendrickson, W. A., Honzatko, R. B. & Sheriff, S. Structural
heterogeneity in protein crystals. Biochemistry 25, 5018–5027 (1986).
541. Wilson, M. A. & Brunger, A. T. The 1.0 A crystal structure of Ca(2+)-bound
calmodulin: an analysis of disorder and implications for functionally relevant
plasticity. J. Mol. Biol. 301, 1237–1256 (2000). 184
542. Martin-Garcia, J. M., Conrad, C. E., Coe, J., Roy-Chowdhury, S. & Fromme, P.
Serial femtosecond crystallography: A revolution in structural biology. Arch.
Biochem. Biophys. 602, 32–47 (2016).
543. van den Bedem, H., Bhabha, G., Yang, K., Wright, P. E. & Fraser, J. S. Automated
identification of functional dynamic contact networks from X-ray crystallography.
Nat. Methods 10, 896–902 (2013).
544. Modelling dynamics in protein crystal structures by ensemble refinement | eLife.
https://elifesciences.org/articles/00311.
545. van Zundert, G. C. P. et al. qFit-ligand Reveals Widespread Conformational
Heterogeneity of Drug-Like Molecules in X-Ray Electron Density Maps. J. Med.
Chem. 61, 11183–11198 (2018).
546. Wall, M. E., Wolff, A. M. & Fraser, J. S. Bringing diffuse X-ray scattering into
focus. Curr. Opin. Struct. Biol. 50, 109–116 (2018).
547. Brereton, A. E. & Karplus, P. A. Native proteins trap high-energy transit
conformations. Science Advances 1, e1501188 (2015).
548. Kabsch, W. Integration, scaling, space-group assignment and post-refinement. Acta
Cryst D 66, 133–144 (2010).
549. Otwinowski, Z. & Minor, W. [20] Processing of X-ray diffraction data collected in
oscillation mode. in Methods in Enzymology vol. 276 307–326 (Academic Press,
1997).
550. Concentric-flow electrokinetic injector enables serial crystallography of ribosome
and photosystem II | Nature Methods. https://www.nature.com/articles/nmeth.3667. 185
551. Sauter, N. K., Hattne, J., Grosse-Kunstleve, R. W. & Echols, N. New Python-based
methods for data processing. Acta Cryst D 69, 1274–1282 (2013).
552. Uervirojnangkoorn, M. et al. Enabling X-ray free electron laser crystallography for
challenging biological systems from a limited number of crystals. eLife 4, e05421
(2015).
553. Adams, P. D. et al. PHENIX: a comprehensive Python-based system for
macromolecular structure solution. Acta Cryst D 66, 213–221 (2010).
554. Schomaker, V. & Trueblood, K. N. On the rigid-body motion of molecules in
crystals. Acta Cryst B 24, 63–76 (1968).
555. Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta
Cryst D 60, 2126–2132 (2004).
556. Davis, I. W. et al. MolProbity: all-atom contacts and structure validation for
proteins and nucleic acids. Nucleic Acids Research 35, W375–W383 (2007).
557. Chapman, H. N., Caleman, C. & Timneanu, N. Diffraction before destruction.
Philosophical Transactions of the Royal Society B: Biological Sciences 369,
20130313 (2014).
558. Toogood, H. S., Leys, D. & Scrutton, N. S. Dynamics driving function − new
insights from electron transferring flavoproteins and partner complexes. The FEBS
Journal 274, 5481–5504 (2007).
559. Catterall, W. A., Wisedchaisri, G. & Zheng, N. The chemical basis for electrical
signaling. Nature Chemical Biology 13, 455–463 (2017).
560. Liu, Y. et al. A pH-gated conformational switch regulates the phosphatase activity
of bifunctional HisKA-family histidine kinases. Nat Commun 8, 2104 (2017). 186
561. Lin, J., Prahlad, J. & Wilson, M. A. Conservation of Oxidative Protein Stabilization
in an Insect Homologue of Parkinsonism-Associated Protein DJ-1. Biochemistry 51,
3799–3807 (2012).
562. Knowles, J. R. Enzyme catalysis: not different, just better. Nature 350, 121–124
(1991).
563. Kamerlin, S. C. L. & Warshel, A. At the dawn of the 21st century: Is dynamics the
missing link for understanding enzyme catalysis? Proteins: Structure, Function, and
Bioinformatics 78, 1339–1375 (2010).
564. Hu, X. et al. The dynamics of single protein molecules is non-equilibrium and self-
similar over thirteen decades in time. Nature Physics 12, 171–174 (2016).
565. García-Meseguer, R., Martí, S., Ruiz-Pernía, J. J., Moliner, V. & Tuñón, I. Studying
the role of protein dynamics in an S N 2 enzyme reaction using free-energy surfaces
and solvent coordinates. Nature Chemistry 5, 566–571 (2013).
566. P. Luk, L. Y., Joel Loveridge, E. & K. Allemann, R. Protein motions and dynamic
effects in enzyme catalysis. Physical Chemistry Chemical Physics 17, 30817–30827
(2015).
567. Cstorer, A. & Ménard, R. [33] Catalytic mechanism in papain family of cysteine
peptidases. in Methods in Enzymology vol. 244 486–500 (Academic Press, 1994).
568. Guan, K. L. & Dixon, J. E. Evidence for protein-tyrosine-phosphatase catalysis
proceeding via a cysteine-phosphate intermediate. J. Biol. Chem. 266, 17026–17030
(1991).
569. Kang, B. S., Devedjiev, Y., Derewenda, U. & Derewenda, Z. S. The PDZ2 Domain
of Syntenin at Ultra-high Resolution: Bridging the Gap Between Macromolecular 187
and Small Molecule Crystallography. Journal of Molecular Biology 338, 483–493
(2004).
570. Kuhn, P. et al. The 0.78 Å Structure of a Serine Protease: Bacillus lentus
Subtilisin,. Biochemistry 37, 13446–13452 (1998).
571. Kelch, B. A., Salimi, N. L. & Agard, D. A. Functional modulation of a protein
folding landscape via side-chain distortion. PNAS 109, 9414–9419 (2012).
572. MacArthur, M. W. & Thornton, J. M. Deviations from Planarity of the Peptide
Bond in Peptides and Proteins. Journal of Molecular Biology 264, 1180–1195
(1996).
573. López-Mirabal, H. R. & Winther, J. R. Redox characteristics of the eukaryotic
cytosol. Biochimica et Biophysica Acta (BBA) - Molecular Cell Research 1783,
629–640 (2008).
574. Sevier, C. S. et al. Modulation of Cellular Disulfide-Bond Formation and the ER
Redox Environment by Feedback Regulation of Ero1. Cell 129, 333–344 (2007).
575. Schierle, C. F. et al. The DsbA Signal Sequence Directs Efficient, Cotranslational
Export of Passenger Proteins to the Escherichia coli Periplasm via the Signal
Recognition Particle Pathway. Journal of Bacteriology 185, 5706–5713 (2003).
576. Santini, C.-L. et al. A novel Sec-independent periplasmic protein translocation
pathway in Escherichia coli. The EMBO Journal 17, 101–112 (1998).
577. TMAO anaerobic respiration in Escherichia coli: involvement of the tor operon -
Méjean - 1994 - Molecular Microbiology - Wiley Online Library.
https://onlinelibrary.wiley.com/doi/abs/10.1111/j.1365-2958.1994.tb00393.x. 188
578. Sauter, N. K. XFEL diffraction: developing processing methods to optimize data
quality. J Synchrotron Rad 22, 239–248 (2015).
579. Zwart, P. H. et al. Automated Structure Solution with the PHENIX Suite. in
Structural Proteomics: High-Throughput Methods (eds. Kobe, B., Guss, M. &
Huber, T.) 419–435 (Humana Press, 2008). doi:10.1007/978-1-60327-058-8_28.
580. Murshudov, G. N. et al. REFMAC5 for the refinement of macromolecular crystal
structures. Acta Cryst D 67, 355–367 (2011).
581. Chen, V. B. et al. MolProbity: all-atom structure validation for macromolecular
crystallography. Acta Cryst D 66, 12–21 (2010).
582. Blakeley, M. P., Hasnain, S. S. & Antonyuk, S. V. Sub-atomic resolution X-ray
crystallography and neutron crystallography: promise, challenges and potential.
IUCrJ 2, 464–474 (2015).
583. Laulumaa, S. & Kursula, P. Sub-Atomic Resolution Crystal Structures Reveal
Conserved Geometric Outliers at Functional Sites. Molecules 24, (2019).
584. The, P. R. et al. Resolution Crystal Structure of a-Lytic. (2004).