Two radical-dependent mechanisms for anaerobic degradation of the globally abundant organosulfur compound dihydroxypropanesulfonate

Article abstract of DOI:10.1073/pnas.2003434117

2(S)-dihydroxypropanesulfonate (DHPS) is a microbial degradation product of 6-deoxy-6-sulfo-d-glucopyranose (sulfoquinovose), a component of plant sulfolipid with an estimated annual production of 1010 tons. DHPS is also at millimolar levels in highly abundant marine phytoplankton. Its degradation and sulfur recycling by microbes, thus, play important roles in the biogeochemical sulfur cycle. However, DHPS degradative pathways in the anaerobic biosphere are not well understood. Here, we report the discovery and characterization of two O2-sensitive glycyl radical enzymes that use distinct mechanisms for DHPS degradation. DHPS-sulfolyase (HpsG) in sulfate- and sulfite-reducing bacteria catalyzes C–S cleavage to release sulfite for use as a terminal electron acceptor in respiration, producing H2S. DHPS-dehydratase (HpfG), in fermenting bacteria, catalyzes C–O cleavage to generate 3-sulfopropionaldehyde, subsequently reduced by the NADH-dependent sulfopropionaldehyde reductase (HpfD). Both enzymes are present in bacteria from diverse environments including human gut, suggesting the contribution of enzymatic radical chemistry to sulfur flux in various anaerobic niches.

Full text of DOI:10.1073/pnas.2003434117

Two radical-dependent mechanisms for anaerobic
degradation of the globally abundant organosulfur
compound dihydroxypropanesulfonate
Jiayi Liua,b,1, Yifeng Weic,1, Lianyun Lina, Lin Tenga, Jinyu Yina, Qiang Lua, Jiawei Chena, Yuchun Zhenga, Yaxin Lia,
Runyao Xua, Weixiang Zhaid, Yangping Liud, Yanhong Liue, Peng Caof, Ee Lui Angc, Huimin Zhaoc,g,2,

Zhiguang Yuchia,2, and Yan Zhanga,b,2
aTianjin Key Laboratory for Modern Drug Delivery & High-Efficiency, Collaborative Innovation Center of Chemical Science and Engineering, School of
Pharmaceutical Science and Technology, Tianjin University, 300072 Tianjin, China; bFrontiers Science Center for Synthetic Biology (Ministry of Education),
Tianjin University, 300072 Tianjin, China; cInstitute of Chemical and Engineering Sciences, Agency for Science, Technology and Research (A*STAR), 138669
Singapore, Singapore; dTianjin Key Laboratory on Technologies Enabling Development of Clinical Therapeutics and Diagnostics, School of Pharmacy, Tianjin
Medical University, 300070 Tianjin, People’s Republic of China; eTechnical Institute of Physics and Chemistry, Chinese Academy of Sciences, 100190 Beijing,
China; fKey Laboratory of Drug Targets and Drug Leads for Degenerative Diseases, Affiliated Hospital of Integrated Traditional Chinese and Western
Medicine, Nanjing University of Chinese Medicine, 210023 Nanjing, China; and gDepartment of Chemical and Biomolecular Engineering, University of Illinois
at Urbana–Champaign, Urbana, IL 61801
Edited by JoAnne Stubbe, Massachusetts Institute of Technology, Cambridge, MA, and approved May 28, 2020 (received for review February 23, 2020)
2(S)-dihydroxypropanesulfonate (DHPS) is a microbial degradation
product of 6-deoxy-6-sulfo-d-glucopyranose (sulfoquinovose), a
component of plant sulfolipid with an estimated annual production
of 1010 tons. DHPS is also at millimolar levels in highly abundant
marine phytoplankton. Its degradation and sulfur recycling by mi-
crobes, thus, play important roles in the biogeochemical sulfur cycle.
However, DHPS degradative pathways in the anaerobic biosphere
are not well understood. Here, we report the discovery and charac-
terization of two O2-sensitive glycyl radical enzymes that use distinct
mechanisms for DHPS degradation. DHPS-sulfolyase (HpsG) in
sulfate- and sulfite-reducing bacteria catalyzes C–S cleavage to
release sulfite for use as a terminal electron acceptor in respiration,
producing H2S. DHPS-dehydratase (HpfG), in fermenting bacteria,
catalyzes C–O cleavage to generate 3-sulfopropionaldehyde, subse-
quently reduced by the NADH-dependent sulfopropionaldehyde re-
ductase (HpfD). Both enzymes are present in bacteria from diverse
environments including human gut, suggesting the contribution of
enzymatic radical chemistry to sulfur flux in various anaerobic niches.
pathway is favored by fermentative bacteria, while the sulfo-ED
pathway is favored by respiratory bacteria (7).
Apart from being a product of sulfoglycolysis, DHPS is also an
important organosulfur compound in its own right. It is present
in up to millimolar intracellular concentrations in eukaryotic
marine phytoplankton, including the highly abundant diatoms (2,
8), which are estimated to contribute ∼20% of the total global
primary production (9). These phytoplankton use sulfate, present
at ∼30 mM levels in seawater to synthesize a variety of organo-
sulfur compounds including DHPS and sulfoquinovose, Secretion
or cell lysis makes these compounds available for degradation by
marine heterotrophic bacteria, accounting for a large component
of the flux of organic carbon in the surface oceans (8, 10).
Significance
DHPS is one of the most abundant organosulfonates on this
planet. The mechanisms for DHPS degradation in the anaerobic
biosphere are not well understood. Here, we report the
bioinformatics-aided discovery, biochemical, and structural char-
acterizations of two O2-sensitive glycyl radical enzymes that use
distinct radical-mediated mechanisms for DHPS degradation in
anaerobic bacteria from diverse terrestrial and marine sources as
well as human gut. These enzymes play an important role in the
biogeochemical sulfur cycle and link dietary sulfonates to micro-
bial production of H2S, which is a causative agent of chronic
diseases, such as inflammation and colorectal cancer.
glycyl radical enzyme sulfoglycolysis dihydroxypropanesulfonate
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sulfate- and sulfite-reducing bacteria gut bacteria
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rganosulfonates are ubiquitous in our environment, our
bodies, and the food that we eat. Mechanisms by which they
O
are metabolized by diverse microbes in the human microbiome
and in the environment are of great relevance to human health
and to the biogeochemical sulfur cycle. Two organosulfonates of
special importance are sulfoquinovose and DHPS due to their
production in large volumes globally by photoautotrophs (1, 2).
Sulfoquinovose is the polar headgroup of the plant sulfolipid
sulfoquinovosyl diacylglycerol (3), a component of photosynthetic
thylakoid membranes in all vascular plants, mosses, algae, and
most photosynthetic bacteria (1). The annual global production of
sulfoquinovose is estimated to be 1010 tons, making it one of the
most abundant organic sulfur compounds in nature (1). Bacterial
degradation of sulfoquinovose was recently discovered and named
sulfoglycolysis due to its resemblance to classical glycolytic path-
ways. The sulfo-Embden–Meyerhof–Parnas (sulfo-EMP) pathway
was characterized in Escherichia coli K-12 and is used during both
aerobic growth (4) and anaerobic mixed acid fermentation (5). In
this pathway, half of the carbon of sulfoquinovose is converted to
dihydroxyacetonephosphate and used for growth, while the other
half is excreted as DHPS, which is, subsequently, metabolized by
other bacteria (5, 6). A second sulfoglycolytic pathway, the sulfo-
Entner–Doudoroff (sulfo-ED) pathway, was characterized in the
environmental isolate Pseudomonas putida SQ1 (7) and produces
3-sulfolactate instead of DHPS. It is thought that the sulfo-EMP
Author contributions: J.L., Y.W., H.Z., Z.Y., and Y. Zhang designed research; J.L., Y.W., L.L.,
L.T., J.Y., Q.L., J.C., Y. Zheng, Y.L., R.X., W.Z., Yanhong Liu, and Y. Zhang performed
research; Yangping Liu, Yanhong Liu, P.C., E.L.A., H.Z., Z.Y., and Y. Zhang contributed
new reagents/analytic tools; J.L., Y.W., L.L., L.T., J.Y., Q.L., J.C., Y. Zheng, Y.L., R.X., W.Z.,
Yanhong Liu, H.Z., Z.Y., and Y. Zhang analyzed data; and J.L., Y.W., E.L.A., H.Z., Z.Y., and
Y. Zhang wrote the paper.
The authors declare no competing interest.
This article is a PNAS Direct Submission.
Published under the PNAS license.
Data deposition: Desulfovibrio vulgaris IseG has been deposited in the Protein Data Bank,
www.wwpdb.org (accession no. 5YMR); two homologs of DvIseG have been deposited in
UniProt, https://www.uniprot.org/ (accession nos. E5Y378 and E5Y7I4); and the crystallog-
raphy, atomic coordinates, and structure factors have been deposited in the RCSB Protein
Data Bank, https://www.rcsb.org (accession no. 6LON).
1J.L. and Y.W. contributed equally to this work.
2To whom correspondence may be addressed. Email: zhao5@illinois.edu, yuchi@tju.
edu.cn, or yan.zhang@tju.edu.cn.
This article contains supporting information online at https://www.pnas.org/lookup/suppl/
doi:10.1073/pnas.2003434117/-/DCSupplemental.
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Various pathways for the degradation of DHPS have been iden-
tified in aerobic environmental bacteria. In Cupriavidus pinatubonensis
(11), (S)-DHPS is racemized in two steps through two DHPS-2-
dehydrogenases (HpsO and HpsP). A DHPS-1-dehydrogenase
(HpsN) then converts (R)-DHPS into (R)-sulfolactate (SL), followed
by C–S cleavage by the enantiospecific (R)-sulfolactate-sulfolyase
(SuyAB) (SI Appendix, Fig. S1A). The sulfite released is oxidized
by the periplasmic sulfite dehydrogenase and excreted as sulfate.
Apart from direct C–S cleavage by SuyAB, two other processes
are known for the desulfonation of (R)-SL. In the aerobic marine
bacterium Roseovarius nubinhibens ISM (12), SL is converted to
cysteate followed by C–S cleavage by cysteate sulfolyase and
converted to sulfoacetaldehyde followed by C–S cleavage by
sulfoacetaldehyde actyltransferase (SI Appendix, Fig. S1A).
In the anaerobic biosphere, which includes environments
ranging from ocean sediments to the digestive tracts of marine
and terrestrial animals, degradation of DHPS by sulfate- and
sulfite-reducing bacteria (SSRB) results in conversion of the
sulfonate sulfur to H2S (5). This process is of particular interest
in the human gut where fermentation of dietary sulfoquinovose
rich in fruits and vegetables produces DHPS, and generation of
H2S by SSRB has been linked to diseases, such as inflammation
and colorectal cancer (13). Despite its importance, pathways for
the degradation of DHPS in anaerobic bacteria are not well
understood. The only organism in which it has been studied is
Desulfovibrio sp. DF1, an isolate from sewage sludge (5). In this
bacterium, (S)-DHPS is imported into the cell through an ABC
Scheme 1. Radical-dependent (S)-DHPS degradation pathways involving
the sulfolyase HpsG and dehydratase HpfG.
the substrate is (S)-DHPS due to the abundance of this organo-
sulfonate in nature.
To test this hypothesis, we recombinantly produced E5Y7I4
transporter and oxidized sequentially to sulfolactate by two NAD+- and its adjacent activating enzyme E5Y7I3 from B. wadsworthia
dependent dehydrogenase DhpA and SlaB (SI Appendix, Fig. S1B).
C–S cleavage by SuyAB releases sulfite, which is used as a terminal confirmed that it was, indeed, a DHPS sulfolyase (Fig. 1), and we
electron acceptor (TEA), producing H2S as an end product.
renamed the GRE and activating enzyme HpsG and HpsH, re-
Previously, we (14) and others (15) reported a radical-dependent spectively. HpsG could be activated by anaerobically recon-
3_1_6 (SI Appendix, Fig. S3). Biochemical characterization
mechanism for C–S cleavage of 2-hydroxyethylsulfonate (isethionate),
which is structurally similar to DHPS, in anaerobic bacteria. The
reaction is catalyzed by the O2-sensitive enzyme IseG, a member
of the glycyl radical enzyme (GRE) superfamily (16–18). GREs
contain an essential O2-sensitive glycyl radical (G•) cofactor,
generated by an activating enzyme through chemistry involving
S-adenosylmethionine (SAM) and a [4Fe–4S]1+ cluster (19, 20),
and catalyze diverse radical-dependent reactions. IseG belongs
to a subset of GREs catalyzing 1,2 lyase reactions, including
C–O, C–N, and C–S lyases (16). The substrates of these lyases
contain a C1–OH group and a variable C2 leaving the group.
Since DHPS contains two OH groups, radical-dependent cleav-
age could, in theory, lead to two possible outcomes: C–S or C–O
cleavage (Scheme 1). Here, we describe our bioinformatics-aided
discovery, biochemical and structural characterizations of two
GREs, a (S)-DHPS sulfolyase (HpsG) and a (S)-DHPS dehy-
dratase (HpfG). HpsG is present in SSRB, while HpfG is present
in fermenting bacteria, and their differing distribution and meta-
bolic roles are discussed.
stituted HpsH (SI Appendix, Fig. S4), forming 0.04 0.02 (out of
a theoretical maximum of 1) (19) G• per dimer (Fig. 1A). The
low radical yield is an issue with many GREs and requires further
analysis. Incubation of activated HpsG with racemic DHPS
resulted in the production of hydroxyacetone (Fig. 1 B–D) and
sulfite as detected using a colorimetric Fuchsin assay (21) (SI
Appendix, Fig. S5A). HpsG exhibited Michaelis–Menten kinetics
for DHPS desulfonation (kcat = 26 1 s−1, Km = 13 2 mM, kcat
/
K
m = 1.9 0.2 mM−1 s−1) (SI Appendix, Fig. S5 B and C) with a
much lower activity for isethionate desulfonation (kcat = 0.57
0.03 s−1, Km = 34 5 mM, kcat/Km = 0.017 0.003 mM−1 s−1
)
(Fig. 1B and SI Appendix, Fig. S5 D and E). (The kcat reported is
normalized by radical content.)
Crystal Structure of HpsG in Complex with (S)-DHPS. To further ex-
amine the origin of the different substrate specificities of HpsG,
we determined the crystal structure of HpsG in complex with
racemic DHPS at 2.2 Å (Fig. 2 and SI Appendix, Fig. S6 and
Table S1). The C121 crystals contain four monomers per asym-
metric unit, each exhibiting a canonical β/α barrel fold common
to other GREs. The overall structure of HpsG is similar to that
of DvIseG with a rmsd of 0.73 Å between 796 Cα atoms (SI
Appendix, Fig. S6A). The well-defined electron density for (S)-
DHPS in the active site of HpsG enabled unambiguous assignment
of the substrate. The C2–OH moiety of the bound (S)-DHPS is
located at a position similar to the C1–OH of isethionate in IseG
and is positioned for deprotonation by Glu464 (Fig. 2A). The
substrate sulfonate group is also coordinated in a similar fashion
as in IseG by Arg183, Gln187, and Arg672 of HpsG. Three water
molecules contribute to the extensive hydrogen bond network
interacting with the substrate (Fig. 2A). However, the orientation
Results
Identification and Biochemical Characterization of the DHPS-Sulfolyase
HpsG. During our previous crystallographic studies of Desulfovibrio
vulgaris IseG (DvIseG, Protein Data Bank [PDB] accession no.:
5YMR) (14), we noted that some close homologs of DvIseG
contain variations in substrate-interacting residues, suggesting the
possibility that they may accept structurally related sulfonate
substrates. We focused on the sulfite-reducing human gut bacte-
rium Bilophila wadsworthia, which encodes two homologs of
DvIseG (UniProt: E5Y378 and E5Y7I4, 60% identity between the
two homologs). Previously, growth of B. wadsworthia RZATAU
with isethionate as the sole TEA resulted in the induction of only of the C3 methylene group of (S)-DHPS is markedly different
E5Y378 but not E5Y7I4 (14). The substrate-interacting residues from that of the corresponding C2 methylene group of isethionate
of DvIseG were identical to those of E5Y378 and were highly with a 160° rotation between the C2–C3–S plane of (S)-DHPS
similar but not identical to those of E5Y7I4 (SI Appendix, Fig. S2), relative to the C1–C2–S plane of isethionate (SI Appendix, Fig.
suggesting a structurally similar substrate. We hypothesized that S6 B and C). The replacement of Phe680 and Thr310 in IseG with
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Fig. 1. EPR spectra and enzymatic assays of HpsG. (A) X-band EPR spectrum of HpsG (40 μM) reconstituted with HpsH (80 μM), SAM (1 mM), and reductant Ti
(III) citrate (100 μM), acquired at a temperature of 90 K and microwave power of 1 mW. (B) Specific activity of HpsG-catalyzed conversion of DHPS to
hydroxyacetone and isethionate to acetaldehyde, measured using NADH-coupled spectrophotometric assays with E. coli glycerol dehydrogenase (GldA) and
Saccharomyces cerevisiae alcohol dehydrogenase (ADH1), respectively. Error bars represent the SD of three individual experiments. (C) Detection of the
products of HpsG-catalyzed reactions by derivatization with 2,4-dinitrophenylhydrazine (DNPH) and liquid chromatography–mass spectrometry (LC–MS)
analysis. (D) Negative ionization mass spectra of DNPH-derivatized reaction products, corresponding to peaks 1 and 2 in C. (Theoretical mass of monoanions,
DNPH hydroxyacetone = 253.0, and DNPH acetaldehyde = 223.0).
the less bulky Ile676 and Ala306 in HpsG accommodates the new stereochemistry is thought to be abstracted in all other mecha-
orientation of the C3 methylene group and the additional C1
hydroxymethyl group of (S)-DHPS (SI Appendix, Fig. S6 B and C).
Binding of DHPS in the active site of HpsG is further established
by a water molecule bridging C1–OH of the substrate and the
hydroxyl group of Ser305 in HpsG, which replaces a Gly309 in
IseG at the same position.
A dual conformation of the thiyl radical residue Cys462 is
observed in HpsG, suggesting high conformational flexibility of
this residue. The minimal distance between the thiyl radial and
the Cα of the glycyl radical residue Gly799 is 4.27 Å (chain D),
and the minimal distance between the C2 of DHPS and the thiyl
radical is 4.09 Å (chain D), which would allow C–H abstraction
to occur (Fig. 2B and SI Appendix, Fig. S6C). In the structure of
DvIseG, the orientation of isethionate suggested that its pro-R
hydrogen is abstracted. The C2–H of (S)-DHPS has the same
orientation, implicating a similar stereochemistry of abstraction
nistically related GREs: CutC, propanediol dehydratase, glycerol
dehydratase, and ribonucleotide reductase (16).
Investigation of DHPS Transporters in SSRB. To identify HpsG in
other organisms, we examined the UniRef50 cluster UniRef50_
A0A348AJ86 (where each member shares ≥50% sequence identity
and ≥80% overlap with the seed sequence of the cluster) (22),
which contains IseG and HpsG sequences. A sequence similarity
network (SSN) was constructed for these proteins, revealing an
organization into two major clusters (Fig. 3A). The first contained
103 proteins, the majority of which have active-site residues iden-
tical to those of IseG from D. vulgaris or D. bizertensis (SI Appendix,
Fig. S2). The second cluster contained 14 proteins with active-site
residues identical to those of B. wadsworthia HpsG (Fig. 3A and SI
Appendix, Table S2) and are, thus, assigned as HpsG.
To find other genes associated with DHPS degradation, the
(SI Appendix, Fig. S6C). In contrast, the hydrogen of the opposite genome neighborhoods of the HpsG sequences were examined
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Fig. S7A) showed enthalpy-driven binding of DHPS with a Kd of
6.7 μM (SI Appendix, Fig. S7B) but no binding of isethionate or
3-hydroxypropane-1-sulfonate (3-HPS) (SI Appendix, Fig. S7 C
and D), confirming that it is a DHPS TRAP transporter
(HpsKLM) (Fig. 3C).
HpsG Is Involved in DHPS Dissimilation in B. wadsworthia. We further
studied the ability B. wadsworthia RZATAU to use DHPS as a
TEA. When pyruvate is supplied as a carbon and electron source,
growth was supported with DHPS as the sole TEA (Fig. 4A),
accompanied by the production of H2S as detected using a methylene
blue assay (Fig. 4A). Sodium dodecyl sulfate polyacrylamide gel
electrophoresis (SDS/PAGE) analysis revealed a prominent pro-
tein band with molecular weight of ∼95 kDa (Fig. 4C), present in
DHPS—but not thiosulfate-grown cells. Mass spectrometric
analysis identified this protein as HpsG (Dataset S3), suggesting
its involvement in DHPS dissimilation in this bacterium.
Identification and Biochemical Characterization of the DHPS Dehydratase
HpfG. Apart from desulfonation, GREs could, in theory, also
catalyze radical-mediated DHPS dehydration, converting it to
3-sulfopropionaldehyde (Scheme 1). Because of the global abun-
dance of (S)-DHPS, we hypothesized the existence of a GRE (S)-
DHPS dehydratase, which would provide an alternative pathway for
DHPS degradation. The catalytic mechanism of GRE diol dehy-
dratases has been extensively studied both experimentally (26) and
computationally (27). The reaction involves five conserved active-
site residues: a Cys thiyl radical involved in substrate radical
generation, Glu involved in substrate C1–OH deprotonation, and
a triad of two His and one Asp involved in coordination and
protonation of the substrate C2–OH leaving group.
To search for a GRE (S)-DHPS dehydratase, 674 candidate
diol dehydratase sequences containing the five conserved active-
site residues were extracted from the 22,000 sequences in the
GRE superfamily (InterPro IPR004184). A SSN was constructed
for these proteins, revealing an organization into two major
clusters (Fig. 5A). The first contained previously characterized
Clostridium butyricum glycerol dehydratase (GDH) and Roseburia
inulinivorans propanediol dehydratase. The second cluster con-
tained 74 GREs of unknown function, corresponding to the UniRef
cluster UniRef50_E3H691 (SI Appendix, Table S3). Examination of
their genome neighborhoods revealed genes that could be involved
in DHPS degradation (Fig. 5B). Like other diol dehydratases, this
GRE is associated with a metal-dependent alcohol dehydrogenase
(ADH), which catalyzes the reduction of the aldehyde product. In
addition, this GRE is associated with a very characteristic
transporter belonging to the TauE family (PF01925), which
includes bacterial sulfite, isethionate, and sulfoacetate exporters,
suggesting a link to sulfonate metabolism. We, therefore, hypothesized
a pathway in which (S)-DHPS is converted to 3-sulfopropionaldehyde
by the GRE (HpfG), reduced to sulfopropanol by the ADH (HpfD),
and excreted through the exporter (HpfE) (Fig. 5C).
Fig. 2. HpsG active-site structure. (A) Structure of the HpsG active site in
complex with the (S)-DHPS (Chain D). The proposed pathway for hydrogen
atom transfer is indicated by red arrows, and hydrogen bonds are indicated
by black dotted lines. 2Fo–Fc electron densities for isethionate are shown at
1.0σ. The two rotamers observed for Cys462 are shown. (B) Proposed
mechanism of DHPS cleavage by HpsG. The thiyl radical, which is transiently
generated by the G• cofactor in all GREs, abstracts a hydrogen (shown in red)
from the substrate DHPS, and returns it to form the product hydroxyacetone.
using the enzyme function initiative–genome neighborhood tool
(Fig. 3B) (23). Two of them (Desulfovibrio litoralis German Collec-
tion of Microorganisms [DSM] 11393, uncultured Desulfovibrio sp
Tax ID: 167968) were associated with transporters in the MFS
family (Pfam PF07690). These were closely related to the Cupriavidus
pinatubonensis (S)-DHPS transporter HpsU (60% sequence identity)
(11), suggesting that they are DHPS transporters (Fig. 3C).
Several of the HpsG homologs, including B. wadsworthia
HpsG, were also associated with a tripartite ATP-independent
periplasmic (TRAP) transporter, a family of transporters that
catalyzes the import of carboxylic and sulfonic acids (24) and
includes the previously characterized isethionate transporter IseKLM
from Desulfovibrio piger (14). TRAP transporters are reliant on a
soluble periplasmic substrate-binding protein (SBP), which is
amenable to biochemical characterization by isothermal titration
calorimetry (ITC) (14, 25). The recombinant SBP (E5Y7I1) of the
HpsG-associated TRAP transporter in B. wadsworthia (SI Appendix,
To test this hypothesis, we recombinantly produced HpfG
(A0A318FL05) and its activating enzyme HpfH (A0A318FEA4)
from Klebsiella oxytoca (SI Appendix, Fig. S8) and HpfD (A0A374P995)
from Hungatella hathewayi (SI Appendix, Fig. S9A). HpfD cata-
lyzed NAD+-dependent oxidation of 3-HPS (SI Appendix, Fig.
S9B) and is presumed to also catalyze the NADH-dependent
sulfopropionaldehyde reduction because of the thermodynamic
reversibility of this reaction. HpfG could be activated by anaero-
bically reconstituted HpfH (SI Appendix, Fig. S1A), forming
0.03 0.01 (out of a theoretical maximum of 1) (19) G• per dimer
(Fig. 6A). The low radical yield is an issue with many GREs, and
requires further analysis. Incubation of activated HpfG with race-
mic DHPS resulted in the production of 3-sulfopropionaldehyde
(Fig. 6 B–D). HpfG exhibited Michaelis–Menten kinetics for DHPS
dehydration (kcat = 130
4 s−1, Km = 5.0
0.6 mM, and
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Fig. 3. SSN and genome neighborhoods of HpsG homologs. (A) SSN for IseG homologs belonging to UniRef50_A0A348AJ86, displayed at the E-value cutoff
of 10−360. Each node represents a unique protein sequence, and the most highly related proteins are grouped together in clusters, putatively sharing the same
function. Blue, active-site residues identical to D. vulgaris or Desulfovibrio bizertensis IseG; red, active-site residues identical to B. wadsworthia HpsG. (B)
Genome neighborhoods of selected homologs in the HpsG cluster. (C) Proposed pathway for DHPS import and HpsG-dependent degradation in B.
wadsworthia.
kcat/Km = 26 3 mM−1 s−1) (SI Appendix, Fig. S10). (The kcat
reported is normalized by radical content.)
sensitive GREs, HpsG and HpfG, described in this study, provide
radical-dependent pathways for DHPS degradation in anaerobic
bacteria and highlight the diversity of O2-sensitive chemistry oc-
curring in anoxic environments.
Homology Model and Molecular Docking of HpfG. A homology
model of HpfG was constructed using a C. butyricum glycerol
dehydratase structure (PDB 1R9D) as a template (37% se-
quence identity between the two proteins), followed by docking
of the DHPS substrate. The position and the pose of (S)-DHPS
in the model is similar to that of glycerol in GDH (SI Appendix,
Fig. S11). The positions of the conserved Cys464, Glu466, and
the triad Asp478, His191, and His308 (SI Appendix, Fig. S11) are
consistent with the conserved mechanism for these diol dehy-
dratases. The substrate sulfonate group interacts with Arg363,
Val677, Tyr481, and Asp478, which are conserved in the mem-
bers of the HpfG cluster. However, given the low sequence
identity between HpfG and the used template, further structural
studies are required and are currently underway.
The HpsG sequences that we have identified are present in
gram-negative SSRB (SI Appendix, Table S2). Like in B. wadsworthia,
the HpsG gene clusters appear to be involved in the uptake of (S)-
DHPS, followed by desulfonation, producing sulfite for use as a
TEA (Fig. 3C). The HpfG sequences are present in gram-negative
Gammaproteobacteria, and gram-positive Clostridia and Lacto-
bacilliales bacteria (SI Appendix, Table S3). Several of the HpfG
sequences in Gammaproteobacteria are associated with homologs
of E. coli sulfoglycolysis enzymes, including sulfoquinovose
isomerase (YihS), sulfofructose kinase (YihV), sulfofructose-1-
phosphate aldolase (YihT), and sulfolactaldehyde reductase
(YihU) (Fig. 5B), suggesting the metabolism of (S)-DHPS produced
internally through sulfoglycolysis (Fig. 5C). HpfG sequences in other
bacteria are associated with a protein annotated as a sodium:sulfate
symporter (PF00939), which we hypothesize is a (S)-DHPS importer,
suggesting the uptake and metabolism of (S)-DHPS from the
environment (Fig. 5C).
Discussion
DHPS is produced in large volumes globally, and the ability of its
two hydroxyl groups to serve as reactive handles for enzymatic
chemistry provides a wide range of possible pathways for DHPS
degradation in metabolically diverse microbes. The two O2-
Many of the HpsGs and HpfGs are from gut bacteria, in-
cluding strains from the human gut microbiome (SI Appendix,
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conduct all anaerobic experiments in an atmosphere of N2 and with the O2
maintained at less than 5 ppm.
Gene Synthesis, Molecular Cloning, and Plasmid Construction. DNA fragments
encoding E. coli codon-optimized sequences of B. wadsworthia HpsG (BwHpsG,
E5Y7I4), HpsH (BwHpsH, E5Y7I3), HpsK (BwHpsK, E5Y7I1), K. oxytoca HpfG
(KoHpfG, A0A318FL05), HpfH (KoHpfH, A0A318FEA4), and H. hathewayi HpfD
(HhHpfD, A0A374P995) were synthesized and inserted into plasmids by General
Biosystems, Inc. (Anhui, China). Plasmids used in this study are pET-28a(+) and
modified pET28 vectors including HMT (containing, in tandem, a His6-tag,
maltose binding protein [MBP], and a tobacco etch virus (TEV) protease
cleavage site, followed by the construct of interest), and HT (containing a
His6-tag and a TEV protease cleavage site, followed by the construct of in-
terest). The BwHpsG, KoHpsG, and HhHpfD fragments were inserted into HT
at the SspI site. For the production of soluble BwHpsH, BwHpsK, and
KoHpfH, the respective fragments were inserted into HMT at the SspI site.
The N-terminal 22-a.a. (amino acid) signal peptide of BwHpsK was omitted.
For the production of high-purity BwHpsG with a N-terminal 20-a.a. trunca-
tion for crystallography, the BwHpsG fragment was amplified by PCR using
primers 1F and 1R (SI Appendix, Table S4) and inserted into the HMT at the
SspI site to form HMT-BwHpsG (-20 a.a.). A surface-entropy reduction mu-
tation, changing a.a. 129–132 from TDMQ to AAAA, was introduced by
QuikChange site-directed mutagenesis using primers 2F and 2R to form
HMT-BwHpsG (-20 a.a. 129TDMQ132-AAAA).
Production of BwHpsGH and KoHpfGH for Enzymatic Assays. BwHpsGH and
KoHpfGH were heterologously expressed in E. coli BL21 (DE3) cells harboring
the plasmids HT-BwHpsG, HMT-BwHpsH and HT-KoHpfG, HMT-KoHpfH, re-
spectively. For BwHpsH, the cells were cotransformed with the plasmid pTf16
(TaKaRa) for coexpression of the tig chaperone. Expression was carried out
in LB supplemented with the appropriate antibiotics (50 μg/mL kanamycin
for BwHpsG, KoHpfG, and MBP-KoHpfH, 50 μg/mL kanamycin and 25 μg/mL
chloramphenicol for MBP-BwHpsH). Starter cultures (5 mL) were inoculated
from single colonies and grown overnight at 37 °C followed by dilution into
1 L of medium. To induce chaperone expression for the MPB-BwHpsH cul-
ture, 2.5 mg/mL L-arabinose was added. The cultures were grown in a
shaking incubator at 37 °C and 220 rpm, up to an optical density at 600 nm
(OD600) of 0.6–0.8. The temperature was then decreased to 18 °C, followed
by addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final con-
centration of 0.4 mM, and continued shaking for 16 h. The cultures were
centrifuged (8,000 × g for 10 min at 4 °C), and the cell pellets (∼1 g wet
weight) were suspended in 5 mL of lysis buffer (50 mM Tris/HCl, pH 8.0,
200 mM KCl, 1 mM phenylmethanesulfonyl fluoride (PMSF), 0.2 mg/mL
lysozyme, 0.03% Triton X-100, and 0.02 mg/mL of DNase I), and frozen in
a −80 °C freezer.
For protein purification, the cells were thawed and incubated at room
temperature (RT, 25 °C) for 50 min, during which cell lysis occurs. Some 5 mM
β-mercaptoethanol (BME) was added, and nucleic acid was removed by
precipitation with 1% streptomycin sulfate. For BwHpsG and KoHpfG, the
protein solution was loaded on to a 5 mL TALON Co2+ column (Clontech
Laboratories, Inc.), preequilibrated with buffer A [20 mM Tris/HCl, pH 7.5,
5 mM BME, and 0.2 M KCl]. The column was washed with 10 column volumes
(CV) of buffer A, and protein was eluted with 5 CV of buffer A containing
150 mM imidazole. The eluted protein was precipitated with solid (NH4)2SO4
to 70% saturation and isolated by centrifugation (20,000 × g for 10 min at
4 °C). The pellet was dissolved in 5 mL of buffer B [20 mM Tris/HCl, pH 7.5,
100 mM KCl, and 5 mM BME] and desalted using a G25 column (GE, ther-
mostat jacket tube XK16/20, packed 15 cm × 2 cm, 30 mL), preequilibrated
with buffer B.
Fig. 4. DHPS-dependent growth of B. wadsworthia and H2S formation. (A)
Various sulfur containing TEAs support the growth of B. wadsworthia. (B)
Formation of methylene blue by reacting the headspace gas with N,N-di-
methyl-pphenylenediamine dihydrochloride and the absorbance of the re-
spective reaction mixtures at 670 nm. The assays were performed in triplicate
and are presented with SDs. (C) B. wadsworthia grown on pyruvate plus
thiosulfate (lane 2), taurine (lane 3), or pyruvate plus DHPS (lane 4). The
arrows indicate a ∼95-kDa band identified as HpsG.
Tables S2 and S3). The production of H2S by human gut SSRB
has been linked to diseases, and HpsG and HpfG may provide
competing pathways for DHPS degradation in metabolically
distinct bacteria. HpsG and HpfG are also present in environ-
mental bacteria, including marine bacteria, and identification of
their sequences allows further investigation of their roles in the
recycling of abundant marine sulfonates (2, 8). HpsG is present
in Desulfovibrio desulfuricans ND132, isolated from mesohaline
Chesapeake Bay sediments and Desulfovibrio sp. HK-II, isolated
from sediments of a partial salt lake (SI Appendix, Table S2).
HpfG is present in many marine bacteria, including gram-negative
Vibrio and Photobacterium species isolated from seawater, sedi-
ments, and marine animals and gram-positive Epulopiscum species
found in fish gut (SI Appendix, Table S3), suggesting a role in
marine anaerobic niches.
In conclusion, our study has expanded the repertoire of en-
zymes and transporters involved in the biodegradation of DHPS,
a highly abundant organosulfonate on this planet. Existing sequence
databases reveal the occurrence of these proteins in microbes
inhabiting diverse anaerobic environments, and their identifica-
tion facilitates further bioinformatics studies of the biological
recycling of this compound.
For BwHpsH and KoHpfH, the protein solution was loaded on to a column
packed with 20 mL amylose resin (New England Biolabs), and protein was
eluted with buffer A containing 10 mM maltose. The purified BwHpsG
(e280 = 122,000 M−1 cm−1), MBP-BwHpsH (e280 = 88,200 M−1 cm−1), KoHpfG
(e280 = 118,000 M−1 cm−1), and MBP-KoHpfH (e280 = 101,000 M−1 cm−1) were
examined by SDS/PAGE on a commercial gel (SurePAGE, Bis-Tris, 8–16%).
Materials and Methods
Materials and General Methods. Lysogeny Broth (LB) was prepared with yeast
extract and tryptone purchased from Oxoid, UK. For LC–MS, high-purity
methanol and acetonitrile from Concord Technology and formic acid from
Merck were used. The ultrapure deionized water used in all experiments was
prepared using a Millipore Direct-Q. Synthetic oligonucleotide primers were
purchased from Tsingke, Inc. (Beijing, China). “ÄKTA pure” or “ÄKTA prime
plus” fast protein liquid chromatography machines were used to perform all
protein purification chromatographic steps. Proteins were quantified by mea-
suring their absorption at 280 nm, using a BioPhotometer D30 (Eppendorf,
Hamburg, Germany). A Lab2000 glovebox (Etelux, Beijing, China) was used to
Production of N-Terminal 20 a.a. Truncated BwHpsG for Protein Crystallization.
Expression of a MBP-BwHpsG mutant for crystallography was carried out in
E. coli BL21 (DE3) cells harboring the plasmid pET28-HMT-BwHpsG (-20 a.a.
129TDMQ132-AAAA). The transformant was grown in LB medium containing
kanamycin (50 μg/mL) at 37 °C in flasks in a shaker incubator at 220 rpm and
induced for the expression of MBP-BwHpsG with 0.3 mM IPTG for 16 h at
18 °C. Typically, cells from 6 L culture were harvested by centrifugation
(8,000 × g for 10 min) and were suspended in 5 mL (∼1 g wet weight) of buffer
C [50 mM Tris/HCl, pH 8.0, 200 mM KCl, 1 mM PMSF, 0.03% Triton X-100]. Then,
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Fig. 5. SSN and genome neighborhoods of HpfG homologs. (A) SSN for putative diol dehydratases in the GRE superfamily, containing conserved residues
thought to be required for catalysis, displayed at the E-value cutoff of 10−260. Each node represents a unique protein sequence, and the most highly related
proteins are grouped together in clusters, putatively sharing the same function. (B) Genome neighborhoods of selected homologs in the HpfG cluster. (C)
Proposed pathways for DHPS import and HpfG-dependent degradation.
the cells were lysed with sonication (Scientz Biotechnology, Ningbo, China). The
lysate was then centrifuged (20,000 × g 30 min at 4 °C) to remove unbroken
cells and cell debris. The protein solution was then applied to a 10 mL TALON
column. The protein was eluted with buffer A containing 150 mM imidazole.
The eluate was then diluted twofold with buffer A and loaded on a column
packed with 40 mL amylose resins. The column was then washed with 5 CV
buffer A and eluted with 5 CV buffer A containing 10 mM maltose. The eluate
premixed with recombinant His6-tagged TEV protease (15:1) was dialyzed
overnight against 2 L buffer A. The dialyzed sample was loaded on a 10 mL
TALON column to retain TEV and MBP proteins. The flow through was col-
lected and dialyzed against 2 L buffer D [20 mM Tris·HCl, pH 7.5, 5 mM BME] for
3 h before it was applied to a 5 mL Q Sepharose high performance column. The
column was eluted with 10 CV linear salt gradient from 100 to 500 mM KCl in
buffer D. A prominent peak containing BwHpsG (-20 a.a.) was collected and
concentrated to a final volume of 5 mL (4 mg/mL) using a centrifuge concen-
trator (30K MWCO; Millipore). This protein solution was then injected into a
Superdex200 gel filtration column (300 mL) and eluted with buffer E [20 mM
Tris·HCl, pH 7.5, 0.2 M KCl, 1 mM dithiothreitol (DTT)]. The eluate from gel
filtration column was reconcentrated and buffer exchanged with the storage
buffer [10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid/KOH, pH 7.4,
50 mM KCl, 1 mM Tris-(2-carboxyethyl)-phosphine hydrochloride]. The final
concentration is 10 mg/mL The purified protein was examined by SDS/PAGE gel.
[Fe-S] Cluster Reconstitution for MBP-HpsH and MBP-HpfH. Reconstitution of
the [Fe–S] clusters of MBP-HpsH and MBP-HpfH, quantitation of Fe and S
contents, and measurement of their absorption spectra, were carried out
according to previously described procedures (14). The purity of MBP-
BwHpsH and MBP-KoHpfH was estimated to be 69% and 73% based on
densitometry analysis of the Coomassie-stained SDS/PAGE gel using the
software ImageJ and used to estimate the Fe and S contents per monomer.
Sequence alignments suggested that HpsH and HpfH, like many other
previously characterized GRE activating enzymes, contain one [4Fe–4S]
cluster in a radical SAM domain, and two [4Fe–4S] clusters in a ferredoxin
domain (SI Appendix, Fig. S12) (28), giving a theoretical maximum of 12 Fe
and 12 S per monomer. Anaerobic reconstitution of the [4Fe–4S] clusters
resulted in 7.9 0.3 Fe and 8.4 0.1 S per HpsH monomer and 5.4 0.2 Fe
and 5.3 0.4 S per HpfH monomer, suggesting that our protocol gives rise to
incomplete reconstitution.
The ultraviolet–visible (UV–Vis) absorption spectra exhibited features
characteristic of [4Fe–4S]2+ clusters, which disappeared upon reduction (SI
Appendix, Fig. S4). Correcting for the 69% purity of MBP-BwHpsH and 73%
purity of MBP-KoHpfH, the extinction coefficients of the reconstituted MBP-
BwHpsH and MBP-KoHpfH [4Fe–4S] clusters were estimated to be 27 and
23 mM−1 cm−1. Assuming an approximate e410nm of 15 mM−1 cm−1 per
cluster (29), we estimate that for MBP-BwHpsH and MBP-KoHpfH, ∼1.8
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Fig. 6. EPR spectra and enzymatic assays. (A) X- band EPR spectrum of HpfG (40 μM) reconstituted with HpfH (80 μM), SAM (1 mM), and reductant Ti (III)
citrate (100 μM), acquired at a temperature of 90 K and microwave power of 1 mW. (B) Specific activity of HpfG-catalyzed conversion of DHPS to
3-sulfopropionaldehyde, measured using a NADH-coupled spectrophotometric assay with sulfopropionaldehyde reductase (HpfD). Error bars represent the SD
of three individual experiments. (C) Detection of HpfG-catalyzed 3-sulfopropionaldehyde formation by derivatization with DNPH and LC–MS analysis. (D)
Negative ionization mass spectra of the DNPH-derivatized reaction product, corresponding to peak 1 c. (Theoretical mass of the monoanion of DNPH
sulfopropionaldehyde = 317.0).
[4Fe–4S] clusters and 1.5 [4Fe–4S] clusters per monomer, respectively, were
out according to previously described procedures (14). A commercial
reconstituted consistent with the measured Fe and S contents.
hydroxyacetone standard (LabNetwork, Wuxi, China) was also prepared.
Electron Paramagnetic Resonance Detection of Glycyl Radical Formation.
Continuous wave X-band electron paramagnetic resonance (EPR) spectros-
copy was used to characterize the glycyl radical. A 200 μL reaction mixture
containing 20 mM Tris, pH 7.5, 0.1 M KCl, 40 μM BwHpsG/KoHpfG, 80 μM
reconstituted MBP-BwHpsH/MBP-KoHpfH, 1 mM SAM, 100 μM Ti(III) citrate
was incubated at RT for 60 min/15 min in the glovebox. Then, 10% glycerol
for HpsG, or 10% D-sorbitol for HpfG was added into the reactions. Samples
were loaded into EPR tubes with 4-mm outer diameter and 8-in. length
(Wilmad Lab-Glass, 734-LPV-7), sealed with a rubber stopper, removed from
the glovebox, and frozen in liquid nitrogen prior to EPR analysis. The ex-
perimental spectra for the glycyl radical were modeled with Bruker Xepr
spin fit to obtain g values, hyperfine coupling constants, and linewidths.
Double integration of the simulated spectra was used to measure spin
concentration (14). For both HpsG and HpfG enzymes, EPR quantitation was
performed with triplicate biological samples. The perpendicular mode
X-band EPR spectra were recorded using a Bruker E500 EPR spectrometer.
Data acquisition was performed with Xepr software (Bruker). The EPR
spectra represent an average of 30 scans and were recorded under the fol-
lowing conditions: temperature, 90 K; center field, 3,370 G; range, 200 G;
microwave power, 1 mW; microwave frequency, 9.43 GHz; modulation
amplitude, 2 mT; modulation frequency, 100 kHz; time constant, 20.48 ms;
conversion time, 40 ms; scan time, 81.92 s; and receiver gain, 23 dB.
Fuchsin Assay for Sulfite Detection. Sulfite was detected using a colorimetric
Fuchsin assay according to previously described procedures (14). BwHpsG
activation was carried out as described for the EPR experiments except that
glycerol was omitted. A 100 μL reaction mixture containing 10 μM activated
BwHpsG, 20 mM DHPS or isethionate was incubated at 30 °C for 1 h in the
glovebox. A negative control omitting substrate was also performed. While
the reaction incubated, stock solution A (0.8 M H2SO4, 0.08% Fuchsin, and
1.6% formaldehyde, mixed 7:2:1) was freshly prepared. A 50 μL portion per
reaction sample was mixed with 950 μL of solution A, incubated for 10 min,
and the UV–Vis absorption spectra were collected.
Spectrophotometric Activity Assay for H. hathewayi HpfD. A 200 μL reaction
mixture containing 20 mM Tris·HCl buffer, pH 9.5, 100 mM KCl, 0.6 ng
HhHpfD, 2 mM NAD+, and 100 mM 3-HPS (Aladdin, Shanghai, China) was
incubated at RT in a 96-well plate, and the absorbance at 340 nm was
monitored using a plate reader (Tecan M200, Männedorf, Switzerland).
Enzymatic Assay for Hydroxyacetone Formation by HpsG. A 200 μL reaction
mixture containing 20 mM Tris·HCl, pH 7.5, 0.1 M KCl, 1 μM activated
BwHpsG, 10 μM E. coli GldA, 0.4 mM NADH, and 50 mM DHPS was incubated
at RT for 20 min in the glovebox. The mixture was removed from the glo-
vebox, and its absorbance at 340 nm was measured using a plate reader
(Tecan M200). Control assays omitting either SAM or BwHpsG were also
performed. To investigate the substrate specificity of BwHpsG, assays
substituting DHPS and GldA with 50 mM isethionate and excess S. cerevisiae
ADH1 were also performed as previously described for IseG (14).
LC–MS/MS Assay for Hydroxyacetone and 3-Sulfopropionaldehyde Detection.
BwHpsG and KoHpfG activation was carried out as described for the EPR
experiments except that glycerol/sorbitol was omitted. For BwHpsG, a 100 μL
reaction mixture containing 10 μM activated BwHpsG, 0.1 mM Ti(III) citrate
and 50 mM DHPS (AA blocks) was incubated at 30 °C for 30 min in the
glovebox. Two negative controls omitting SAM or DHPS were also per-
formed. Isethionate (Adamas) added instead of DHPS was also performed.
For KoHpfG, a 100 μL reaction mixture containing 10 μM activated KoHpfG,
0.1 mM Ti(III) citrate, and 20 mM DHPS was incubated at 30 °C for 30 min in
the glovebox. Three negative controls omitting KoHpfG, SAM, or DHPS were
also performed. Derivatization with DNPH and LC–MS analysis were carried
Coupled Spectrophotometric Activity Assays for HpsG. A 100 μL reaction
mixture containing 20 mM Tris·HCl, pH 7.5, 0.1 M KCl, and typically, 1 μM
activated BwHpsG, 10 μM GldA, 0.2 mM NADH, and 100 mM DHPS was in-
cubated at RT in a 1 cm Eppendorf cuvette in the glovebox. The absorbance
at 340 nm was monitored at 4 s intervals using the cuvette mode of a Thermo
Scientific Nanodrop OneC in the glovebox. To obtain the Michaelis–Menten
kinetic parameters, DHPS concentration was varied while keeping a fixed
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enzyme concentration of 1 μM BwHpsG. Assays substituting DHPS and GldA
with isethionate and ADH1 were also prepared, and the absorbance 340 nm
was monitored at 15 s intervals. These assays contained, typically, a saturating
substrate concentration of 500 mM isethionateand 5 μM BwHpsG. The kinetic
data presented were obtained from a HpsG sample with 0.06 G• per protein
dimer. GraphPad Prism6 was used for data analysis.
Growth of B. wadsworthia with Different TEAs. B. wadsworthia strain RZATAU
(DSM 11045) was purchased from DSMZ. Cells were first inoculated into ABB
medium supplemented with 5 mM taurine and cultivated anaerobically at
30 °C for 3–7 d. Then, 100 μL portions of the starter culture were transferred
into three anaerobic bottles containing 5 mL modified DSM 503 medium,
omitting taurine, and supplemented with 60 mM Na-formate and 200 μg/ L
1,4-naphthochinone. Different TEAs (20 mM) were added: 1) Na2S2O3 (with
20 mM sodium pyruvate was added as a carbon and electron source), 2)
taurine, and 3) DHPS (with 20 mM sodium pyruvate was added as a carbon
and electron source). After 3–7 d incubation at 30 °C, all three cultures be-
came turbid and contained a black precipitate. H2S in the headspace gas was
detected using a methylene blue assay as previously described (14). The
samples were diluted prior to absorbance measurement to ensure that they
fall within the linear region of the spectrometer.
Enzymatic Assay for 3-Sulfopropionaldehyde Formation by HpfG. A 100 μL
reaction mixture containing 20 mM Tris·HCl, pH 7.5, 0.1 M KCl, 0.5 μM ac-
tivated KoHpfG, 10 μM sulfopropionaldehyde reductase (HhHpfD), 0.2 mM
NADH, and 100 mM DHPS was incubated at RT for 2 min in the glovebox.
Additionally, then its absorbance at 340 nm was measured using the cuvette
mode of a Nanophotometer NP80 Mobile in the glovebox. Control assays
omitting either DHPS or KoHpfG were also performed.
Protein Identification by SDS/PAGE and Mass Spectrometry. Cells were har-
vested by centrifugation, lysed by boiling in Laemmli loading buffer, and
analyzed on a 10% SDS/PAGE gel. Prominent protein bands induced by
growth on sulfonate substrates were manually excised. After in-gel digestion
and extraction, the peptide mixtures were loaded onto Orbitrap Fusion
LUMOS MS. The MS/MS spectra from each LC–MS/MS run were searched
against the B. wadsworthia protein database GCF_000185705.2 (Bilo_-
wads_3_1_6_V2) from UniProt (release date of March 19, 2014; 68,406 en-
tries) using an in-house Proteome Discoverer (Version PD1.4, Thermo-Fisher
Scientific). Protein identifications were performed based on Sequest HT (34).
Source data underlying Fig. 4C are provided as in Dataset S1.
Coupled Spectrophotometric Activity Assays for HpfG. A 100 μL reaction
mixture containing 20 mM Tris·HCl, pH 7.5, 0.1 M KCl, and, typically, 0.5 μM
activated KoHpfG, 10 μM HpfD, 0.2 mM NADH, and 100 mM DHPS was in-
cubated at RT in a 1 cm Eppendorf cuvette in the glovebox. The absorbance
at 340 nm was monitored at 5 s intervals using the cuvette mode of a
Nanophotometer NP80 Mobile in the glovebox. To obtain the Michaelis–
Menten kinetic parameters, DHPS concentration was varied while keeping a
fixed enzyme concentration of 500 nM KoHpfG. The kinetic data presented
were obtained from
a HpfG sample with 0.04 G• per protein dimer.
GraphPad Prism6 was used for data analysis.
X-ray Crystal Structure of BwHpsG. Initial screening of BwHpsG crystals was
performed using an automated liquid handling robotic system (Gryphon, Art
Robbins) in 96-well format by the sitting-drop vapor diffusion method. The
screens were set up at 295 K using various sparse matrix crystal screening kits
from Hampton Research and Molecular Dimensions. Several crystallization
conditions gave thin plate-shape crystals. After further optimization using
the hanging-drop vapor-diffusion method in 24-well plates, we obtained
crystals large enough for single crystal X-ray diffraction studies. The best
condition yielding large platelike crystals was 0.2 M sodium acetate and
0.1 M Bis–Tris propane, pH 6.5, 16% (wt/vol) PEG3350 plus 500 mM DHPS.
Crystals were flash cooled in liquid nitrogen using reservoir solution
containing 25% glycerol as a cryoprotectant. Diffraction data were collected
on a local Rigaku X-ray diffractor (XtaLAB P200 MM007HF, Tokyo, Japan) to
a resolution of 2.20 Å. The data set was indexed, integrated, and scaled
using HKL3000 suite (30). Molecular replacement was performed by PHENIX
(31) using the crystal structure of IseG (PDB 5YMR). The structure was
manually built according to the modified experimental electron density
using Coot (32) and further refined by PHENIX (31) in iterative cycles before
it was deposited in the RCSB Protein Data Bank (accession no. 6LON). The SI
Appendix, Table S1 contains the statistics for data collection and final re-
finement. All structural figures were generated with UCSF Chimera (33).
ITC Assays for BwHpsK. ITC experiments were carried out on a 50 μM solution
of BwHpsK in buffer containing 20 mM Tris·HCl, pH 7.5 and 200 mM KCl with
1 mM DTT using a PEAQ-ITC instrument (Malvern). The samples were titrated
with 19 injections of 2 μL of 1 mM DHPS, isethionate, or 3-HPS at 25 °C and a
stirring speed of 750 rpm. Deionized water was used for the reference cell.
Background titrations were obtained using protein-free buffer, and sub-
tracted from the raw titrations. The data were fitted to a single-site binding
model using the MicroCal PEAQ-ITC software to estimate the stoichiometry,
binding affinity, and changes in enthalpy (ΔH) and entropy (ΔS). A Kd of 6.7
μM was measured for DHPS with a N value of 2.2. No interaction was ob-
served for isethionate and 3-HPS.
Data Availability. Desulfovibrio vulgaris IseG deposited in the Protein Data
Bank, www.wwpdb.org (accession no. 5YMR), two homologs of DvIseG in
UniProt, https://www.uniprot.org/ (accession nos. E5Y378 and E5Y7I4), and
coordinates and structure factors of HpsG in complex with (S)-DHPS have
been deposited in the RCSB Protein Data Bank, https://www.rcsb.org (ac-
cession no. 6LON). All other data required to evaluate the paper’s conclu-
sions are present in the paper and/or the SI Appendix.
ACKNOWLEDGMENTS. We thank the instrument analytical center of School
of Pharmaceutical Science and Technology at Tianjin University for providing
the LC–MS analysis, Zhi Li, and Drs. Xinghua Jin, Yan Gao, and Xiangyang
Zhang for helpful discussion. This work was supported by the National Key
R&D Program of China (Grant 2019YFA0905700), the National Natural Science
Foundation of China (Grant 31870049) (Y. Zhang), National Key R&D Program
of China (Grant 2017YFD0201400, Grant 2017YFD0201403), National Natural
Science Foundation of China (Grant NSFC31972287), the State Key Laboratory
of Ecological Pest Control for Fujian and Taiwan Crops (Grant SKL2019001,
Grant SKL2019003) (Z.Y.), and the Agency for Science, Research and Technol-
ogy of Singapore Visiting Investigator Program Grant 1535j00137 (H.Z.).
Homology Modeling and Docking. A homology model of HpfG was created by
Prime module of Schrödinger software (Schrödinger LLC, New York, NY)
using C. butyricum glycerol dehydratase structure (PDB 1R9D) as a template.
The sequence identity between the HpfG and the template protein is ∼37%.
The ligand (S)-DHPS was sketched in Chem-DrawUltra 13.0 and, sub-
sequently, prepared using the LigPrep module of Schrödinger software. (S)-
DHPS was docked by Glide module of Schrödinger using an extra precision
mode to a reference position in the binding site of HpfG similar as glycerol in
the template structure (PDB 1R9D).
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