MftD Catalyzes the Formation of a Biologically Active Redox Center in the Biosynthesis of the Ribosomally Synthesized and Post-translationally Modified Redox Cofactor Mycofactocin

Article abstract of DOI:10.1021/jacs.9b06102

Mycofactocin (MFT) is a putative ribosomally synthesized and post-translationally modified (RiPP) redox cofactor. The biosynthesis of MFT is encoded by the gene cluster mftABCDEF. While processing of the precursor peptide by MftB, MftC, and MftE has been shown to result in the formation of the small molecule 3-amino-5-[(p-hydroxyphenyl)methyl]-4,4-dimethyl-2-pyrrolidinone (AHDP), no activity has been shown for the putative dehydrogenase MftD and the putative glycosyltransferase MftF. In addition, evidence demonstrating that MFT is a redox cofactor has only been limited to the requirement of mft genes for ethanol assimilation in Mycobacterium smegmatis mc²155. Here, we demonstrate that MftD catalyzes the oxidative deamination of AHDP, forming an α-keto moiety on the resulting molecule, which we call pre-mycofactocin (PMFT). We characterize PMFT by 1D and 2D NMR spectroscopy techniques and by high-resolution mass spectrometry data to solve its structure. We further characterized PMFT by cyclic voltammetry and found its midpoint potential to be ～255 mV. Lastly, we demonstrate that PMFT is a biologically active redox cofactor that oxidizes NADH bound by M. smegmatis carveol dehydrogenase (MsCDH) and can be used by MsCDH in the oxidation of carveol. These data demonstrate for the first time that PMFT functions as a biologically active redox mediator and provides the most direct evidence to date that MFT is a RiPP-derived redox cofactor.

Full text of DOI:10.1021/jacs.9b06102

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Article
MftD Catalyzes the Formation of a Biologically Active Redox
Center in the Biosynthesis of the Ribosomally Synthesized and
Post-translationally Modified Redox Cofactor, Mycofactocin.
Richard S. Ayikpoe, and John A. Latham
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.9b06102 • Publication Date (Web): 05 Aug 2019
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MftD Catalyzes the Formation of a Biologically Active Redox Center
in the Biosynthesis of the Ribosomally Synthesized and Post-transla-
tionally Modified Redox Cofactor, Mycofactocin.
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Richard S. Ayikpoe1 and John A. Latham1
*
1Department of Chemistry and Biochemistry, University of Denver, Denver, Colorado 80210, United States
KEYWORDS: mycofactocin, ribosomally synthesized and post-translationally modified peptide, redox cofactor, oxidative deamina-
tion, carveol dehydrogenase, MftD
ABSTRACT:
Mycofactocin (MFT) is a putative ribosomally synthesized and post-translationally modified (RiPP) redox cofactor. The bio-
synthesis of MFT is encoded by the gene cluster mftABCDEF. While processing of the precursor peptide by MftB, MftC, and MftE has been
shown to result in the formation of the small molecule 3-amino-5-[(p-hydroxyphenyl)methyl]-4,4-dimethyl-2-pyrrolidinone (AHDP), no ac-
tivity has been shown for the putative dehydrogenase MftD and the putative glycosyltransferase MftF. In addition, evidence demonstrating
that MFT is a redox cofactor has only been limited to the requirement of mft genes for ethanol assimilation in M. smegmatis mc2155. Here, we
demonstrate that MftD catalyzes the oxidative deamination of AHDP, forming an a-keto moiety on the resulting molecule which we call pre-
mycofactocin (PMFT). We characterize PMFT by 1D and 2D nuclear magnetic resonance spectroscopy techniques and by high-resolution
mass spectrometry data to solve its structure. We further characterized PMFT by cyclic voltammetry and found its midpoint potential to be
~255 mV. Lastly, we demonstrate that PMFT is a biologically active redox cofactor that oxidizes NADH bound by M. smegmatis carveol
dehydrogenase (MsCDH) and can be used by MsCDH in the oxidation of carveol. These data demonstrate for the first time that PMFT
functions as a biologically active redox mediator and provides the most direct evidence to date that MFT is a RiPP-derived redox cofactor.
INTRODUCTION
the MFT associated dehydrogenase families were shown to contain
non-exchangeable nicotinamide cofactors that were active in vitro
Ribosomally synthesized and post-translationally modified pep-
tides (RiPPs)have emerged as a structurally and biologically diverse
class of secondary metabolites.1 Interest in RiPPs is particularly fo-
cused on the immense bioavailability and engineerability of diverse
antibiotics.2–5 While the vast majority of characterized RiPPs have
been shown to possess antimicrobial activity, others have been
shown to have more diverse biological functions such as quorum
sensing and redox activity.6–13 Until recently, this latter activity was
defined by the sole quintessential member, pyrroloquinoline qui-
none (PQQ), which is used as an electron acceptor in bacterial sugar
and alcohol dehydrogenases.11,14,15 However, it was proposed that
the uncharacterized mycofactocin biosynthetic pathway could en-
code for the second member of the RiPP-derived redox cofactor
family.16 While, the mycofactocin biosynthetic pathway bears a
striking resemblance to that of PQQ, the bona fide structure and
function of the molecule remains unknown.
A
mftR
mftB
mftD
mftF
mftA
mftC
mftE
SDR
MDRETEAETAELVTESLVEEVSIDGMCGVY
MftA**
B
MftA
MftC
dAdo
SAM
SAM
dAdo
MftC
Mycofactocin (MFT) was first identified by a bioinformatic anal-
ysis that demonstrated the cooccurrence of the genes mftABCDEF
(Figure 1A)16 in over 600 bacterial species, including a score of hu-
man pathogens (eg. M. tuberculosis, M. avium, and M. ulcerans).16,17
In addition, the same bioinformatic study indicated that genes en-
coding for three different dehydrogenase families (TIGR03971,
TIGR03989, and TIGR04266) were found in the genome only
when mft genes were present. This codependence upon the mft bi-
osynthetic pathway implicates an important role for MFT in the ac-
tivities of the aforementioned TIGR families. Notably, members of
MftE
MftA1-28 H O
MftD
2
AHDP
MftA*
Figure 1
– Mycofactocin biosynthetic pathway. A) An arrow repre-
sentation of the mycofactocin biosynthetic pathway showing the as-
sociation of a short chain dehydrogenase (SDR) and the MuMftA
amino acid sequence. B) A condensed reaction scheme of known
steps in mycofactocin biosynthesis. Enzymatic modifications are
shown in red.
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A
B
0.2
C
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D
FMN
FA
Riboflavin
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25 kDa
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300
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Retention Time (min)
Wavelength (nm)
Figure 2
– A) MftD was purified to homogeneity as determined by SDS-PAGE analysis. Lane 1 contains the Goldbio BLUEstain Protein
Ladder standard, Lane 2 contains His6-purified MuMftD, and lane 3 contains His6-purified MsMftD. B) A UV-Vis spectral analysis of anaer-
obically prepared MftD indicates the presence of a reduced flavin (red). Upon exposure to air of the protein sample, the UV-Vis spectrum
shifts considerably, indicating that the flavin is oxidized. C) High pressure liquid chromatography analysis of the flavin containing fraction of
protein precipitate (red) indicates that MftD likely binds FMN when compared to retention times of flavin standards.
with only non-physiologically relevant redox mediators.18,19 This fur-
ther suggested that MFT could be required for catalytic turnover for
these enzymes. In support of this notion, recent gene knockout stud-
ies have indicated that mftA-D and mftF, along with the iron-de-
pendent alcohol dehydrogenase msmeg_6242 (TIGR04266), are
essential for ethanol assimilation in the model organism M. smegma-
tis mc2155.20 In addition, the same study demonstrated that disrup-
tion of the mft genes led to imbalance of cellular concentrations of
NAD+/NADH, indicating for the first time in vivo, that MFT may
play a role in redox metabolism. However, direct evidence for the
physiological role of MFT has not been provided, in part because its
structure remains unknown.
present a combination of activity assays, nuclear magnetic resonance
studies, and high resolution-mass spectrometry studies to demon-
strate conclusively that MftD catalyzes the oxidation of the L-amino
moiety on AHDP to form the redox center of MFT. Moreover, we
provide compelling evidence that the resulting molecule is redox ac-
tive and is shown to serve, in vitro, as a catalytically competent co-
factor for the putative MFT-dependent carveol dehydrogenase,
MsCDH. Together, our experimental data provides strong evidence
that supports the notion that MFT is a novel, catalytically-compe-
tent, RiPP-derived, redox cofactor.
RESULTS
Recent work by our lab, and others, has focused on elucidating
the in vitro biosynthesis, and thereby the structure and function, of
MFT. It was proposed that MFT is synthesized from the conserved
C-terminal region on precursor peptide MftA.16 Validation of this
proposal was provided when two independent studies reported that
MftC catalyzes the S-adenosylmethionine dependent oxidative de-
carboxylation of the C-terminus on MftA, in the presence of MftB,
resulting in an a/b unsaturated tyramine (Figure 1B, MftA**).21,22
However, upon further investigation it was shown that MftC cataly-
sis also resulted in the a functionally relevant conversion of MftA**
to MftA* which contains a bicyclic modification consisting of a 5-
membered lactam ring derived from the penultimate valine residue
(Figure 1B).23 These results led to the examination of MftE catalysis
where it was shown that the enzyme specifically hydrolyzes MftA*,
resulting in the formation of 3-amino-5-[(p-hydroxyphenyl)me-
thyl]-4,4-dimethyl-2-pyrrolidinone (AHDP, Figure 1B).24 The re-
maining gene products, MftD and MftF, encode for putative lactate
dehydrogenase and glycosyl transferase respectively, and their func-
tion in mycofactocin biosynthesis have been hitherto unknown.
MftD is an FMN Binding Protein.
The mftD gene (mul_0774)
was cloned from M. ulcerans Agy99 and the MuMftD protein was
heterologously expressed in and anaerobically purified from Esche-
richia coli (Figure 2A). Characterization of as-isolated MuMftD
protein by UV-visible absorbance spectroscopy indicated the
presences of a reduced flavin which is characterized by the broad
shoulder between 300 nm - 410 nm (Figure 2B, red). Following ex-
posure to air, the absorbance spectra of MuMftD shifted to that of
an oxidized flavin, characterized by the dual absorbance maxima at
360 nm and 445 nm (Figure 2B, blue). To determine which species
of flavin was bound, MuMftD was heat precipitated and the soluble
fraction was analyzed by HPLC. The retention time of the flavin in
the soluble fraction was compared to authentic flavin standards (fla-
vin adenine dinucleotide, FMN, and riboflavin, Figure 2C). From
this analysis, it was evident that the absorbance features of the cofac-
tor bound to MuMftD originated from FMN, consistent with other
members of the protein-fold family.32 Notably, when MuMftD was
purified aerobically, the protein appeared to lose FMN during the
isolation process. Consistent with this observation, analysis of
MuMftD by analytic HPLC-SEC demonstrated that upon oxida-
tion, the protein releases FMN (Figure S1), in the absence of sub-
strate. In our hands, we observed that the loss of FMN is irreversible.
MftD belongs to the aldolase-TIM barrel fold family that is com-
prised of a-hydroxy acid dehydrogenases which are known to be fla-
vin mononucleotide (FMN) dependent.25–27 This putative FMN de-
pendency led us to speculate that MftD could catalyze the molecular
oxygen dependent oxidation of the phenyl ring of AHDP to generate
a the corresponding catechol, reminiscent to PQQ and other redox
cofactors.28–31 The rationale for this proposal was that if MFT was
redox active, it required a physiologically relevant redox center
which was not apparent on the AHDP parent molecule. However,
without any evidence, this proposal was only speculative. Herein, we
Reconstitution of MftD Activity.
Our expectation was that
AHDP, the product from the MftE reaction, might be the substrate
for MftD. To validate this hypothesis, reactions containing AHDP
and MftD were carried out aerobically and the reaction mixture was
analyzed by HPLC. The aerobic addition of AHDP to MftD and
subsequent analysis by HPLC does in fact give rise to a new species
at a retention time of ~15.8 min (Figure 3A, blue) with concomitant
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Structural Elucidation of the MftD Product.
disappearance of AHDP (~14.1 min); the associated kobs for the sin-
To validate that
gle turnover reaction was measured to be 0.010 ± 0.002 min-1 (Fig-
ure S2A). Interestingly, the UV-visible absorbance spectrum for the
new species is substantially different than that of AHDP, indicating
that the resulting modification also impacted the electronics of the
molecule (Figure S2B and S2C). To ensure that this activity was not
unique to MuMftD, the gene msmeg_1424, encoding for MsMftD,
was cloned from M. smegmatis mc2155, the protein was purified
from E. coli (Figure 2A), and reactions with AHDP were prepared
and analyzed by HPLC. As expected, the HPLC chromatogram for
the MsMftD reaction showed similar results to that of MuMftD
(Figure 3A, red), indicating that AHDP is most likely the substrate
for both Ms and MuMftD. Herein, we worked solely with MuMftD
and will refer to the protein as simply MftD.
1
MftD catalyzes the oxidative deamination of AHDP and the subse-
quent formation of PMFT, 1D and 2D NMR studies were carried
out on the isolated product. Reactions containing AHDP and MftD
were performed on a large scale (~2 mg AHDP) and the HPLC pu-
rified PMFT was analyzed by NMR. For 13C NMR, we used synthe-
sized MftA labeled with U-13C valine and U-13C tyrosine at positions
Val29 and Tyr30 in reactions with MftB, MftC, MftE, and MftD to
generate the U-13C labeled PMFT. This was required to increase the
13C NMR signal due to the limited quantities of isolated PMFT. Alt-
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hough we had already reported on the H, 13C, and COSY NMR
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characterization of AHDP, we further analyzed it by HSQC 2D
NMR in this study. This allowed us to directly compare AHDP
spectra to that of PMFT.
Overlaid COSY spectra of the lactam region for both AHDP (red)
and PMFT (blue) are shown in Figure 4A. Notably, the chemical
shifts of the protons within the lactam are modestly shifted down-
field by 0.1-0.4 ppm. This downfield shift is accompanied by the dis-
appearance of the H a in the 1H NMR spectrum for PMFT, suggest-
ing that Ca was the site of modification. Consistent with this obser-
vation, 13C NMR indicated a drastic downfield shift of the Ca in
AHDP from ~61 ppm to ~210 ppm in PMFT (Figure 4B and see
Supplementary Information for full spectra). The chemical shift of
Cais corroborated by HMBC spectra (Figure 4B) which shows long
range interaction between the protons on the geminal methyl and
the Ca. Lastly, overlaid HSQC spectra of AHDP and PMFT show
the loss the proton-carbon coupling associated with H-Ca. Together
these NMR data provide strong evidence that MftD catalyzes the ox-
idative deamination of AHDP to form PMFT.
A
B
234.1107 235.1445
AHDP
+ MsMftD
+ MuMftD
13
14
15
16
17
233
234
235
236
237
Retention Time (min)
m/z
[M+H+]
235.1445 m/z
[M+H+]
234.1125 m/z
C
MftD
Figure 3
– A) HPLC chromatograms of reactions containing
AHDP (black), AHDP and MsMftD (red), and AHDP and
MuMftD (blue) indicate that AHDP is an active substrate for MftD.
B) HRMS analysis of the AHDP (black) and the MftD product
(blue) shows an ion with a m/z that is consistent with the loss of -
NH3 and the addition of O. C) From the HPLC and HRMS anal-
ysis, MftD is proposed to catalyze the oxidative deamination of
AHDP (black) to form pre-mycofactocin (blue, PMFT). The the-
oretical m/z for the molecules are indicated above their structure.
A
Hα
B
Hδ
2.1
2.3
2.5
2.7
2.9
3.1
3.3
3.5
3.7
3.9
4.1
10
30
50
70
90
110
130
150
170
190
210
Cα
1.15
1.05
To determine the nature of the MftD catalyzed modification, we
analyzed the product of the reaction by high resolution mass spec-
trometry (HRMS). Consistent with our previous work,24 HRMS
analysis of the AHDP [M+H]+ ion, reanalyzed here, was found to
have a mass-to-charge ratio (m/z) = 235.1445 (Figure 3B, black),
within 4 ppm of the predicted mass (C13H18N2O2, theoretical m/z =
235.1441 Da). HRMS analysis of the ~15.8 min MftD product pro-
vided a m/z = 234.1107 (Figure 3B, blue). This mass is consistent
with the loss of NH3 and the addition of a single oxygen atom to
ADHP (C13H15NO3, theoretical [M+H]+ m/z = 234.1125). These
results are inconsistent with our previous proposal suggesting that
MftD could catalyze the formation of a catechol on the phenyl ring
of AHDP. Instead, our HRMS analysis suggests that MftD might
catalyze the oxidative deamination of AHDP, installing an a-keto
moiety, resulting in the formation of 5-[(p-hydroxyphenyl)methyl]-
4,4-dimethyl-2,3-pyrrolidinedione (Figure 3C, blue) or herein re-
ferred to as pre-mycofactocin (PMFT). Precedence for flavin de-
pendent oxidative deamination comes from D-amino acid oxidase
(DAAO). DAAO’s function to catalyze the flavin adenine dinucleo-
tide (FAD) dependent and stereospecific oxidation of D-amino acid
to form a-keto acids.33,34 However, it should be noted that DAAO
and MftD belong to different protein fold families.35,36
4.1
3.5
2.3 2.1
3.9 3.7
3.3 3.1 2.9 2.7 2.5
Hα
C
D
E
δ
δ
30
40
50
60
β
α
Cα
4.0 3.8 3.6 3.4 3.2 3.0 2.8 2.6 2.4 2.2
Figure 4
– NMR analysis of PMFT. A) Overlaid COSY spectra of
AHDP (red) and PMFT (blue) reveals the loss of the Ha signal.
B) HMBC spectra for PMFT provides evidence that the Ca has
shifted downfield to ~210 ppm. C) Overlaid HSQC spectra of
ADHP (red) and PMFT (blue) shows the loss of the H-Ca hetero-
nuclear interaction. D) A reference structure of PMFT with the
important carbon annotated. E) A summary of relevant NMR cor-
relations on PMFT.
The Role of Oxygen in the MftD Reaction
PMFT structure, we next sought to determine the source of the
Having solved the
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oxidase prior to the addition of MftD. Glucose oxidase catalyzes the
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oxidation of glucose to gluconate and hydrogen peroxide and is of-
ten added to reactions to achieve anaerobicity.38,39 The subsequent
HPLC chromatogram of the anaerobic reaction indicated only the
presence of the starting material AHDP (Figure 5B). This suggests
that molecular oxygen is required for the MftD activity. Since anaer-
obically purified MftD contains FMNH2, and since we did not ob-
serve any turnover of MftD, it is likely that O2 is required for the ox-
idation of FMNH2. Notably, MftD was inactive in reactions carried
out under the same conditions but with the addition of NAD+ (Fig-
ure S4) suggesting that the nicotinamide cannot fulfill the role of O2.
While efforts to measure the existence and stoichiometry of the pu-
tative peroxide were not undertaken, the results from this analysis is
consistent with O2 being the final electron acceptor in MftD cataly-
sis.
Scheme 1
idase (DAAO) and B) for MftD.
– A) A condensed reaction scheme for D-amino acid ox-
incorporated oxygen atom. Since MftD catalyzed a similar reaction
to DAAO, we turned to its well-characterized mechanism for guid-
ance. The DAAO reaction is described by the two-step oxidative de-
amination of the D-amino group on the amino acid (Scheme 1A).
To begin with, DAAO oxidizes the amino acid to an a-imimo acid
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through a hydride transfer from the Ca to FAD.37 This is followed
by a nucleophilic substitution by an activated water and the subse-
quent oxidation to form the ketone.33 We rationalized that MftD
could carry out a similar reaction mechanism which we describe in
Scheme 1B. To determine if water was the source of the inserted ox-
ygen, we carried out MftD reactions in 99% enriched 18OH2 and an-
alyzed the resulting PMFT by HRMS. From this experiment, we ex-
pected to observe an enrichment of a PMFT ion that was consistent
with a single 18O being incorporated (+2 m/z). As expected, the
mass spectrum of the reaction mixture carried out in 18OH2 shows
two predominant ions (Figure 5A), one at the expected m/z for un-
labeled PMFT ([M+H]+ m/z = 234.1125) and the other at a m/z =
236.1169, within 2 ppm of the expected mass for the 18O-incorpo-
rated PMFT ([M+H]+ m/z = 236.1167). The percent incorpora-
tion was ~65%, near the expected theoretical maximum of 87%.
Conversely, when the reaction was carried out in 18O2, we did not
observe significant incorporation of 18O in the product (Figure
S3A). The rate of back exchange was determined to not significantly
affect the overall incorporation of 18O during the course of this ex-
periment (Figure S3B). Therefore, these results are consistent with
water being the source of the inserted oxygen atom in the MftD cat-
alyzed oxidative deamination of AHDP.
PMFT is Redox and Functionally Active In Vitro.
As noted
previously, the structure of AHDP did not have an apparent physio-
logically relevant redox center. However, following oxidative deam-
ination by MftD, we rationalized that the resulting a-keto-amide
moiety could be the active site of MFT. Indeed a-keto acids are a
common 2e-/2H+ redox moiety and can be found in biologically im-
portant molecules such as pyruvate, a-ketoglurate, and oxaloacetate.
To provide insight about the possible redox behavior of PMFT, cy-
clic voltammetry was used to directly measure the midpoint poten-
tial of PMFT. Three-electrode CV experiments were carried out us-
ing a glassy carbon working electrode, an Ag/AgCl reference elec-
trode, and a platinum counter electrode. PMFT was non-covalently
adsorbed onto the working electrode using single walled carbon
A
B
5×102
1.5
1.0
0.5
0.0
0 µM
[PMFT]
0
-5×102
C
150 µM
-0.50 -0.25 0.00
0.25
0.50
250
300
350
400
450
[M+H+]
236.1167 m/z
Potential (V) vs SHE
Wavelength (nm)
D
[M+H+]
A
B
234.1125 m/z
236.1152
16O/18OH2
AHDP
234.1113
-O2
14 15 16 17 18 19 20 21
Retention Time (min)
233 234 235 236 237 238
m/z
+O2
E
13
14
15
16
17
234
235
236
Retention Time (min)
m/z
Figure 5
– A) High resolution mass spectra of the MftD reaction
carried out in 18OH2 shows the enrichment of the 18O incorporated
PMFT (red). The structure for natural abundance and enriched
PMFT are shown above with their theoretical mass for the [M+H]+
ion. B) HPLC analysis of MftD reactions under anaerobic (blue)
and aerobic (red) conditions suggest that molecular oxygen is re-
quired for catalytic turnover. The reference chromatogram for
AHDP is shown in black.
Figure 6
– A) Overlaid cyclic voltammogram of PMFT/SWCNT
(red) and buffer/SWCNT (black). Voltammetry of PMFT was
measured at pH 7.0 and at 22 oC with a scan rate of 50 mV/s. B)
Overlaid UV-visible spectra showing the oxidation of MsCDH
bound NADH by PMFT. C) HPLC analysis demonstrates that
MsCDH is active towards carveol in the presence of PMFT (blue).
The chromatogram for PMFT is shown in red. D) HRMS analysis
of the ~14.9 min peak shows an ion with a m/z that is consistent
with the mass of PMFTH2 (theoretical [M+H]+ m/z = 236.1281).
E) The proposed structure of PMFT following a 2e-/2H+ reduction.
In the DAAO mechanism and the proposed mechanism for MftD,
O2 is the final electron acceptor.33 To verify that molecular oxygen
is required for MftD turnover, reactions were carried out anaerobi-
cally. To achieve anoxic conditions, reactions were conducted in an
anaerobic chamber and supplemented with glucose and glucose
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the absorbance of DCPIP at 600 nm (DeDCPIP 600nm = 20.7 mM-1cm-1,
nanotubes (SWCNT) as a co-absorbent. CV measurements yielded
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a single signal that was observed between -100 mV and -400 mV as
compared to the buffer/SWCNT control (Figure 6A). From this
measurement, the reductive and oxidative potentials were found to
be -370 mV and -140 mV respectively, and the overall midpoint po-
tential to be -255 mV. We did not observe significantly differenti-
ated peaks for two single electron transfer events, thus we define the
midpoint potential for the 2e-/2H+ transfer as -255 mV.
Figure S8A) due to spectral overlap with NADH. These reactions
were fitted to a linear equation which provided a kobs < 0.01 s-1, sub-
stantially slower than that observed for PMFT. Control stopped-
flow experiments were carried out to ensure that DCPIP was not ox-
idizing MsCDH on the millisecond timescale (Figure S8B). While
there is a slim possibility that DCPIP could oxidize MsCDH faster
than the deadtime of the instrument (3 ms), the more likely scenario
is that DCPIP is non-specifically interacting with MsCDH which
leads to the slow rate of oxidation. Overall, the >100-fold rate en-
hancement of PMFT over DCPIP further indicates that (P)MFT is
a physiological reductant of MsCDH.
Having shown that PMFT is redox active, we next postulated that
it could be used to oxidize NADH in MFT-dependent dehydrogen-
ases. M. smegmatis mc2155 carveol dehydrogenase (MsCDH,
Msmeg_1410) was chosen as the model MFT-dependent dehydro-
genase to test this hypothesis since the R. erythropolis DCL14 hom-
olog has been shown to catalyze the oxidation of carveol to carvone
using 2,6-dichlorophenolindophenol (DCPIP) as an electron accep-
tor19 and since it was known to copurify with NADH.18 This latter
property is significant because it could allow us to monitor the
PMFT dependent oxidation of NADH to NAD+ by monitoring UV-
visible absorbance at 340 nm (DeNADH 340nm = 6.22 mM-1cm-1). To
demonstrate that PMFT could be used as an oxidant, scanning UV-
visible spectroscopic measurements were carried out with purified
MsCDH (see Figure S5 for an SDS-PAGE). As shown in Figure 6B,
the incremental addition of PMFT resulted in a stoichiometric de-
crease in absorbance at 340 nm suggesting that PMFT is mediating
the oxidation of NADH to NAD+ on MsCDH. It should be noted
that we did not observe the same oxidative activity towards free
NADH (Figure S6A). Consequently, this implies that PMFT is bi-
ologically active with MsCDH and potentially with other MFT de-
pendent dehydrogenases.
After finding that PMFT was capable of oxidizing NADH bound
to an MFT-dependent dehydrogenase, we next sought to couple
PMFT to the MsCDH catalyzed oxidation of carveol. To do so,
MsCDH was incubated with excess amounts of carveol and PMFT
in a stoichiometric ratio. The subsequent reaction mixture was an-
alyzed by HPLC (Figure 6C). We observed a change in the reten-
tion time of PMFT in the chromatogram, from ~15.8 to ~14.9 min,
with an accompanying change in the UV-visible spectrum (Figure
S7A and S7B). In addition, we observed a concomitant appearance
of new species at a retention time of ~19.5 min corresponding to car-
vone (for a full HPLC analysis see Figure S6B). We expected that
the species at ~14.9 min is the reduced form of PMFT (PMFTH2).
To provide evidence for this hypothesis, we analyzed the HPLC pu-
rified species by HRMS. As expected, HRMS analysis (Figure 6D)
of the ~14.9 min species provided a m/z = 236.1267 which is con-
sistent with the m/z of PMFTH2 (C13H17NO3, theoretical [M+H]+
m/z = 236.1281). Taken together, the observation that PMFT oxi-
dizes the NADH on MsCDH, PMFT facilitates MsCDH carveol ox-
idation activity, and the observation of a mass consistent with
PMFTH2, provides compelling evidence that PMFT is the biologi-
cally relevant redox moiety of MFT.
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1.1
1.0
0.9
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Time (s)
– Stopped-flow kinetic analysis of a single turnover ox-
Figure 7
idation reaction with 160 µM MsCDH and 25 µM (red dashes)
or 50 µM (magenta dashes) PMFT. The oxidation of NADH at
340 nm was monitored during the reaction, sampling every 2
ms. Each kinetic trace is an average of 3 individual experiments.
The averages were fitted to a single exponential decay (black) to
determine the rate constant.
DISCUSSION
Nature has used ribosomally synthesized and post-translationally
modified peptides (RiPPs) to produce a diverse array of bioactive
molecules. In particular, the use of a RiPP biosynthetic pathway to
produce a small molecule, such as pyrroloquinoline quinone
(PQQ), seems gratuitous considering the high energetic costs asso-
ciated with the synthesis of the precursor peptide. In the case of
PQQ, of the ~30 amino acids on the precursor peptide PqqA, only
tyrosine and glutamate are conserved in the final compound.40–42
Likewise, in the case of MFT biosynthesis only two residues from a
~30 amino acid peptide are being used to make MFT. This implies
that the function of the molecules contributes more to the physiol-
ogy of the cell than the energetic cost to make them. As it happens,
this is the case for PQQ, which serves to transfer electrons from
sugar and alcohol dehydrogenases to the electron transport chain,43
and thus participates in the generation of cellular currency, ATP. By
analogy, the same could be predicted for MFT.
To determine the efficacy of PMFT as a redox mediator, the ap-
parent rate constant (kobs) governing the oxidation of MsCDH
bound NADH by PMFT was determined. To do so, time-depend-
ent stopped-flow UV-vis spectrometry experiments were carried out
at two PMFT concentrations, in triplicate each, monitoring the oxi-
dation of MsCDH bound NADH at 340 nm (Figure 7). Data were
fitted to a single exponential decay which yielded a kobs = 0.8 ± 0.1 s-
1. To directly compare this rate to the rate of oxidation of MsCDH
by DCPIP, we carried out similar stopped-flow reactions monitoring
Nearly a decade ago it was proposed that MFT is the second
member of the RiPP-derived redox cofactor family.16 The assertion
of this proposal was solely based on a bioinformatic analysis that
concluded that associated dehydrogenases might require MFT for
catalytic turnover and that the MFT biosynthetic pathway appeared
reminiscent to that of PQQ. Over a short period of time, we have
seen the maturation of MFT take place. The discovery that MftC
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catalyzed a two-step cyclization to from the bicyclic MftA* provided
functioning as a biologically active redox cofactor. However, when
intrigue to the possibility that MFT could be a redox cofactor.21–23
The demonstration of the subsequent hydrolysis of MftA* to form
the small molecule AHDP further fueled this hypothesis.24,44 Lastly
and significantly, recent work has shown that mft genes are required
for the in vivo ethanol assimilation in M. smegmatis mc2155.20 Alt-
hough, these findings have coalesced to suggest that MFT is a redox
cofactor, no direct evidence has been provided to date.
compared to the remaining peptide-derived redox cofactors a star-
tling and significant structural difference is observed: PMFT is not a
quinone like its brethren. Instead the structure of PMFT consists of
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an a-keto amide moiety which can be reduced by a 2e-/2H+ process,
similar to that of a quinone. Indeed, not only is the functionality of
the a-keto amide moiety similar to the quinones but its electro-
chemical properties are too. The PMFT midpoint potential (-255
In this work, we have provided evidence that AHDP is an active
substrate for MftD. The NMR structural workup of the MftD prod-
uct, paired with HRMS analysis, provides conclusive evidence that
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MftD catalyzes the oxidative deamination of AHDP to form the a-
keto moiety on PMFT. The existence of the a-keto moiety on
PMFT led us to postulate that the molecule could be redox active.
To probe for redox activity, we used cyclic voltammetry to measure
the midpoint potential of PMFT. The observance of redox activity
suggested to us that PMFT is likely the work horse of the MFT mol-
ecule, much like F0 is for F420 and riboflavin is for FMN and FAD.
Consequently, we considered that PMFT might be recruited and
harnessed by MFT-dependent dehydrogenases and therefore we
probed for coupled enzymatic reactivity with the putative carveol de-
hydrogenase MsCDH. Accordingly, PMFT was shown to cycle
MsCDH and was shown to be reduced to PMFTH2, clearly indicat-
ing that PMFT undergoes a 2e-/2H+ reduction by MsCDH. More
importantly, this provides compelling evidence that PMFT, and
thereby MFT, is a biologically active redox cofactor.
Figure 8.
The structures of the known peptide derived redox co-
factors where TPQ, LTQ, TTQ, and CTQ are formed in situ of
the active enzyme and PQQ and PMFT are formed from a dedi-
cated RiPP biosynthetic pathway.
The discovery of novel FMN-dependent AHDP deaminase activ-
ity for MftD is extraordinary. To begin with, it continues to validate
our previous findings that MftC is responsible for the two-step mod-
ification of MftA and that MftE hydrolyzes MftA* to form AHDP.
Although this could be regarded as superficial, the fact that the elu-
cidation of MFT biosynthesis has been successfully executed with-
out any prior structural knowledge of MFT is remarkable. Valida-
tion of MFT intermediates (eg. AHDP) has only come from the sub-
sequent enzyme (eg. MftD) recognizing the intermediate as a sub-
strate. In addition to validating previous work, the reconstitution of
MftD also pulls in parallels with that of PQQ biosynthesis. In PQQ
and MFT biosynthesis, the first biosynthetic step requires a radical-
S-adenosylmethionine protein dependent intramolecular C-C bond
formation.23,41 In both pathways this step is followed by the subse-
quent excision of the crosslinked moiety by a protease24 and the oxi-
dation of the resulting small molecule.42,45 The similarities between
PQQ and MFT biosynthesis was first identified by Haft through bi-
oinformatics and that analysis appears to have withstood experi-
mental scrutiny.16 It would be worthwhile to use the combined PQQ
and MFT biosynthetic pathway architectures as the basis for the dis-
covery of potentially new RiPP derived redox cofactors or interest-
ing small molecules.
Although only RiPP derived redox cofactors have been discussed
thus far, it should be noted that other non-RiPP peptide derived re-
dox cofactors are known. The four known examples are trihydroxy-
phenylalanine quinone (TPQ), tryptophan tryptophylquinone
(TTQ), lysyl tyrosine quinone (LTQ) and cysteine tryptophylqui-
none (CTQ) (Figure 8).14 Although PQQ does belong to the pep-
tide-derived redox cofactor family, there are substantial differences
to the remaining quinones. Whereas PQQ is synthesized through
dedicated biosynthetic pathways, the aforementioned quinones are
synthesized in situ of the functional enzyme.29–31 Here, we propose
that MFT should be permanently added to the list of peptide derived
redox cofactors. As previously mentioned, PMFT is capable of
mV) is comparable to that of the remaining quinones which have
measured potentials in the range of -150 mV to -240 mV.46–48
In terms of functionality, we show here that PMFT, and by exten-
sion MFT, is likely the physiologically relevant cofactor for MsCDH.
It is interesting to note that mft containing genomes also encode a
wide variety and quantity of the MFT-associated dehydrogenases.16
For instance, the M. smegmatis mc2 155 genome encodes for eight
members of the TIGR03971 family and four members of the
TIGR03989 family. The work here provides the only in vitro and
direct evidence that MFT is biologically active with MsCDH, a
member of the TIGR03971 family. Of the remainder MFT-
dependent dehydrogenases, Msmeg_6242, a putative primary alco-
hol dehydrogenase, is the only other dehydrogenase in M. smegma-
tis that has a potential activity associated with it.20 Beyond these two
examples and their respective homologues, it could be that MFT
participates in many biological processes.
Our studies herein now raise the question: What is the final my-
cofactocin structure? The remaining enzyme in the pathway is the
putative glycosyltransferase, MftF. However, the role it is playing in
mycofactocin biosynthesis is, more now than ever, convoluted.
Considering that PMFT is recognized by at least one MFT-
dehydrogenase, it could be that MftF is being used to decorate the
p-hydroxy of the phenyl ring. Alternatively, MftF could be used in
another unknown function unrelated to MFT biosynthesis and that
PMFT could represent the final MFT structure. In summary, we
demonstrate for the first time that MftD catalyzes the oxidative de-
amination of AHDP, forming the redox active center in PMFT.
Moreover, we measured the midpoint potential for PMFT and
demonstrated that it is biologically active with the carveol dehydro-
genase MsCDH. Taken together, we provide the most direct evi-
dence that MFT is the second member of the RiPP derived redox
cofactor family.
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centrifuged at 14000 rpm for 10 min to remove all precipitated pro-
METHODS
1
teins. The supernatant was pipetted into a 1.5 mL HPLC au-
tosampler vial and analyzed by reverse-phase chromatography on a
Shimadzu Prominence-i LC-2030C HPLC using a Jupiter C 18, 5
µm, 4.6 x 250-mm column (Phenomenex), and 10 mM sodium
phosphate monobasic, pH 5.5 (Buffer A) and 90% methanol (Buffer
B) as the mobile phase. A linear gradient of solvent B from 0% to
35% was applied from 2 to 7 min, which was followed by another
linear gradient to 95% solvent B from 7 to 15 min. A third gradient
of solvent B from 95% to 100% was applied from 15 to 17 min after
which solvent B was held constant at 100% from 17 to 19 min. This
was followed by linear decrease to 100% solvent A from 19 to 22 min.
Solvent A was held constant for an additional 2 min to re-equilibrate
the column before subsequent sample injections. Chromatograms
were reported at 450 nm while monitoring wavelengths between
200 and 600 nm. Authentic samples of riboflavin, flavin mononucle-
otide (FMN) and flavin adenine dinucleotide (FAD) were run as
controls.
Cloning, Expression, and Purification of MftD from Mycobacterium
ulcerans Agy99. The mul_0774 gene sequence encoding for MftD
from Mycobacterium ulcerans Agy99 (Uniprot: 0PM50) was
cloned into pET28a-TEV using BamHI and XhoI restriction sites.
The sequence verified MumftD/pET28a-TEV plasmid was trans-
formed into E. coli ArcticExpress (DE3) (Stratagene) for protein
production. An overnight culture of the E. coli ArcticExpress (DE3)-
MumftD/pET28a-TEV grown in 50 mL of terrific broth (TB) me-
dium, was used to inoculate 4 L of the same medium containing 50
µg/mL of kanamycin and 10 µg/mL of tetracycline and supple-
mented with 100 µM riboflavin. The culture was incubated at 37 oC
with shaking at 200 rpm until an OD600 ~ 0.6 was attained, at which
point MftD production was induced by addition of IPTG to a final
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o
concentration of 1 mM. Following a 24 h induction at 13 C with
shaking at 200 rpm, the cells were harvested by centrifugation at
7000 rpm for 10 min and stored at -80 OC until lysis. All purification
steps were carried out in Coy Lab anaerobic chamber maintained
under an atmosphere of 97% N2 and 3% H2 gases. The harvested
cells were thawed and resuspended in five times pellet mass of lysis
buffer containing 2X phosphate buffered saline (PBS) buffer (pH
8.0), 30 mM imidazole and 2 mM dithionite (DTH). To the suspen-
sion was added 0.1 mg/g of lysozyme, 0.1 mg/g of DNase and 1 %
w/v CHAPS and stirred on ice for an additional 30 min. Cells were
disrupted by sonication for 5 min to complete the lysis. The resulting
lysate was clarified by centrifugation at 13,000 rpm for 10 min and
the supernatant was loaded onto a 5 ml HisTrap FF Ni-NTA column
(GE Healthcare) pre-equilibrated with lysis buffer using an AKTA
Start FPLC (GE Healthcare). The column was washed with 2X PBS
buffer (pH 8.0) containing 50 mM imidazole and the bound protein
was eluted using 2X PBS buffer (pH 8.0) containing 300 mM imid-
azole and 2 mM DTH. Fractions containing MftD proteins were im-
mediately buffer exchanged into 2X PBS storage buffer (pH 8.0)
containing 10% glycerol and 2 mM DTH using PD-10 columns (GE
Healthcare). The resulting volume containing MftD protein was
concentrated using 30 kDa spin concentrators (Millipore). To-
bacco-etch virus (TEV) protease was added to the protein and incu-
bated at room temperature for 3 h to hydrolyze the N-terminal His-
tag. The protein/protease mixture was loaded onto a 5 mL HisTrap
FF Ni-NTA column pre-equilibrated with storage buffer. The flow
through was collected and the bound protein was eluted with 2X
PBS (pH 8.0) containing 50 mM imidazole and 2 mM DTH. Frac-
tions containing MftD proteins were pooled together, buffer ex-
changed into a fresh 2X PBS storage buffer (pH 8.0) containing 10%
glycerol using PD-10 columns (GE Healthcare). The resulting pro-
tein fraction was concentrated using 30 kDa spin concentrators and
subjected to a final size exclusion chromatography purification step
as described below.
AHDP Modification Reactions. AHDP modification reactions were
conducted in volumes of 200 µL under aerobic and anaerobic condi-
tions and contained 50 mM sodium phosphate (pH 8.0) as the reac-
tion buffer. Consecutively, 100 µM MftD, 200 µM FMN and 100 µM
of AHDP (see Supplementary Information for isolation procedures)
were added and the reactions were incubated under aerobic and
anerobic conditions for 12 h. Control reactions were set up in a sim-
ilar fashion except buffer was substituted for individual reagents,
protein or substrate. Reactions were then filtered through 0.2 µm
spin columns to remove all precipitated proteins. The supernatant
was pipetted into 300 µL autosampler HPLC vials and injected di-
rectly onto a reverse-phase Jupiter C5, 5 µm, 4.6 x 250-mm column
(Phenomenex) using Shimadzu Prominence-i LC-2030C HPLC
and 5 mM sodium phosphate, pH 7.5 (Buffer A) and 5 mM sodium
phosphate in 70% acetonitrile pH 7.5 (Buffer B). A linear gradient
of solvent B from 0% to 35% was applied from 2 to 8 min, which was
followed by a linear gradient to 100% solvent B from 8 to 17 min.
Solvent B was then held constant at 100% from 17 to 19 min fol-
lowed by linear decrease to 100% solvent A from 19 to 22 min. Sol-
vent A was held constant for an additional 2 min to re-equilibrate
the column before subsequent sample injections. Chromatograms
were reported at 280 nm while monitoring wavelengths between
200 and 400 nm. New peaks were isolated, lyophilized and analyzed
by NMR and HRMS.
Oxygen Requirement for MftD Reaction with AHDP. To investigate
the dependence of MftD catalysis on oxygen, a total of 200 µL of
MftD reaction was set up under anaerobic conditions as follows. To
the reaction buffer containing 2X PBS (pH 8.0) were added consec-
utively10 mM glucose and 10 µM of glucose oxidase (GOX) fol-
lowed by incubation at room temperature for 10 min to deplete ox-
ygen from the buffer. Sequentially, 100 µM of MftD, 200 µM of
FMN, and 200 µM of AHDP were added to the reaction followed by
an additional 12 h of incubation period at room temperature. Reac-
tions were filtered through 0.2 µm spin columns to remove all pre-
cipitated proteins. The supernatant was pipetted into 300 µL au-
tosampler HPLC vials and injected directly onto a reverse-phase
HPLC system. HPLC analysis was carried out as described above.
Cloning, Expression and Purification of MftD from Mycobacterium
smegmatis. The msmeg_1424 gene encoding for MftD from Myco-
bacterium smegmatis mc2155 (Uniprot: A0QSB) was cloned into
pET28a vector using NdeI and XhoI restriction sites. The sequence
verified msmeg_1424/pET28a plasmid was transformed into E. coli
Arctic Express (DE3) for protein expression. MsMftD proteins were
expressed and purified as described above for MuMftD with throm-
bin being substituted for TEV.
18O Isotope Labelling of PMFT. To determine the source of oxygen
atom incorporated into the product of MftD catalysis, reactions were
carried out in either 18OH2 or 18O2. For reactions carried out in
Flavin Determination in MftD. To determine the type of flavin con-
tained in MftD, 100 µL of 200 µM of the purified protein was first
denatured by heating at 80 oC for 10 min. The denatured protein was
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18OH2, a buffer solution of 25 mM ammonium acetate was first pre-
containing 50 µg/ml of kanamycin. The overnight culture was used
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pared by lyophilizing 500 µl of the buffer solution to powder and re-
suspending into an equal volume of 18OH2 (PET grade, Sigma Al-
drich). Reactions consisted of 90 µl of 18OH2 buffer, 100 µM of MftD
(5 µL), 200 µM of FMN (1 µL) and 100 µM of AHDP (4 µL). Fol-
lowing a 1 h aerobic incubation at room temperature, the reaction
was analyzed by HRMS by direct infusion onto a Thermo Scientific
Q Exactive. Reactions carried out in the presence of 18O2 proceeded
similarly but were perform in 16OH2 buffer that was bubbled with
18O2 for 5-min prior to the addition of MftD. A control reaction was
performed to monitor the back-exchange of labeled oxygen. To do
so, 16O-PMFT was incubated in 18OH2-buffer for 60 min prior to
analysis by HRMS.
to inoculate 4 L of LB growth medium containing 50 µg/ml of kana-
mycin. The culture was incubated at 37 oC with shaking at 200 rpm
until an OD600 of ~0.6, at which point MsCDH production was in-
duced by addition of IPTG to a final concentration of 1 mM and al-
lowed to grow overnight at 20 oC with shaking at 200 rpm. The cells
were harvested by centrifugation at 7000 rpm for 10 min and the pel-
let was resuspended in five times pellet mass of lysis 50 mM MOPS
buffer (pH 7.0) containing, 250 mM NaCl and 30 mM imidazole.
To the suspension was added 0.1 mg/g of lysozyme, 0.1 mg/g of
DNase and 1 % w/v CHAPS and stirred for 30 min. Cells were dis-
rupted by sonication for 5 min on ice to complete the lysis. The re-
sulting lysate was clarified by centrifugation at 13,000 rpm for 10 min
the supernatant was loaded onto a 5 mL HisTrap HP Ni-NTA col-
umn pre-equilibrated with lysis buffer using an AKTA Pure FPLC.
The column was washed with 50 mM MOPS buffer (pH 7.0) con-
taining 250 mM NaCl and 30 mM imidazole and the bound protein
was eluted using 50 mM MOPS buffer (pH 7.0) containing 250 mM
NaCl and 300 mM imidazole. Fractions containing MsCDH pro-
teins were pooled together and immediately buffer exchanged into
50 mM MOPS storage buffer (pH 7.0) containing 250 mM NaCl
and 10% glycerol using a HiPrep 26/10 desalting column. The re-
sulting volume of protein was concentrated using 10 kDa spin con-
centrators. Thrombin protease was added to the concentrated pro-
tein and incubated at room temperature for 1 h to cleave off the N-
terminal His-tag. The protein/protease mixture was loaded onto a
Superdex 200 10/300 GL analytical size exclusion chromatographic
column using an AKTA Pure FPLC. The mobile phase was storage
buffer containing 50 mM MOPS (pH 7.0), 250 mM NaCl, and 10%
glycerol at a flow rate of 1.0 mL/min. Protein peaks were collected
and concentrated using 10 kDa spin concentrators. The concen-
trated protein was aliquoted into 1.5 mL vials, flash-frozen in liquid
nitrogen and stored at -80 oC until use.
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Nuclear Magnetic Resonance Spectroscopy. All NMR spectra were
recorded at the at the University of Denver on a Bruker UltraShield
500/54 Plus spectrometer. All Spectra were processed and analyzed
using TopSpin v. 2.1 program (Bruker). All peptide NMR samples
were prepared in 99.96% deuterated water. Water Suppression sig-
nal was applied at a frequency of 2353.37 Hz for all 1H NMR spectra.
Signals were integrated, and the coupling constants were calculated
in MestReNova v. 10.0.1 program (Mestrelab Research).
High Resolution Mass Spectrometry. All samples were analyzed at
Anschutz Medical Campus by the Biological Mass Spectrometry Fa-
cility at the University of Colorado. 18O-labelled samples were in-
fused directly while all other samples were desalted using a C18
ZipTip (EMD Millipore) and subjected to LC-ESI-MS using a
Thermo Scientific Q Exactive and a nanoflow liquid chromatog-
raphy system. The data was analyzed using XCalibur Qual Browser
v. 3.0.63 (Thermo Scientific).
Electrochemical Characterization of PMFT. To determine the re-
dox potential of PMFT, a 5 mg/ml suspension of functionalized sin-
gle walled carbon nanotubes (SWCNTs) was made by dispersing 50
mg of the nanotubes in 10 ml of dimethylformamide and sonicating
in ultrasonic bath for 1 h to obtain a uniform suspension. Glassy car-
bon electrodes were polished with 0.05 µm alumina on nylon and
microbroth polishing pads. The electrode was sonicated in 10 ml of
water in ultrasonic bath for 5 min followed by 10 ml of ethanol for
additional 5 min and allowed to thoroughly dry at room tempera-
ture. The dried electrode was coated with 20 µL of the SWCNTs sus-
pension and allowed to dry at room temperature. This was repeated
two more times and allowing for thorough drying in between casts.
The dried SWCNTs-modified glassy carbon electrode was incu-
bated in phosphate buffer (pH 7.0) as control or in phosphate buffer
(pH 7.0) containing 500 µM of PMFT for 3 h prior to running cyclic
voltammetry (CV) experiments. A conventional thre-electrode sys-
tem was used in a water-jacketed glass cell. The counter electrode
was platinum wire and Ag/AgCl was the reference electrode. CV
measurements were carried out in Coy Lab anaerobic chamber using
CHI 600E Series Electrochemical analyzer. Measurements were rec-
orded at 0.05 V/s and a potential sweep window of +0.2 to -0.8 V at
22 oC.
Monitoring MsCDH UV Vis Spectral Changes in the Presence of
PMFT. To determine if PMFT could be used as a cofactor by
MsCDH for carveol modification, a UV spectroscopic assay was per-
formed using Shimadzu TCC-240A UV-visible spectroscopy as fol-
lows. An initial UV absorbance spectrum of 200 µL reaction buffer
containing 50 mM MOPS (pH 7.0), 250 mM NaCl, 10% glycerol
and 150 µM MsCDH was taken from 190 nm to 500 nm. Following
initial scan, 30 µM PMFT was added to MsCDH, incubated for 5
min, and the UV-visible spectra was measured. This was repeated
until a final concentration of 150 µM PMFT was achieved. A control
experiment was carried out using the same procedures but with
NADH free in solution.
MsCDH-Carveol Modification Reactions. Carveol modification re-
action was conducted in a total volume of 200 µL containing 50 mM
MOPS buffer (pH 7.0), 250 µM NaCl, 10% glycerol, 100 µM
MsCDH, 500 µM carveol, and 500 µM PMFT. The reaction was in-
cubated at room temperature for an hour. Control reactions were set
up in a similar fashion except buffer was substituted for individual
reagents, protein or substrate. Reactions were filtered through 3 kDa
spin columns to remove all precipitated proteins. The supernatant
was pipetted into 300 µL autosampler HPLC vials and injected di-
rectly onto a reverse-phase HPLC system. The HPLC analysis was
run on a Shimadzu Prominence-i LC-2030C HPLC using a Jupiter
C5, 5 µm, 4.6 x 250-mm column (Phenomenex), and 5 mM sodium
phosphate, pH 7.5 (Buffer A) and 5 mM sodium phosphate in 70%
acetonitrile pH 7.5 (Buffer B). A linear gradient of solvent B from
Cloning, Expression, and Purification of Carveol Dehydrogenase
(CDH). The msmeg_1410 gene sequence encoding CDH from
Mycobacterium smegmatis mc2155 (Uniport: A0QSA5) was cloned
into pET28a using NdeI and HindIII restriction sites. The sequence
verified msmeg_1410/pET28a plasmid was transformed into the E.
o
coli BL21 star (DE3) and grown overnight at 37 C in 50 mL LB
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0% to 35% was applied from 2 to 8 min, which was followed by a phosphate buffer saline; PMFT, premycofactocin; PQQ, pyrrolo-
1
2
3
4
5
6
linear gradient to 100% solvent B from 8 to 17 min. Solvent B was
then held constant at 100% from 17 to 19 min followed by linear de-
crease to 100% solvent A from 19 to 22 min. Solvent A was held con-
stant for an additional 2 min to re-equilibrate the column before
subsequent sample injections. UV absorbance readings were moni-
tored between 200 nm and 400 nm.
quinoline quinone; SAM, S-adenosylmethionine; SEC, size exclu-
sion chromatography; SHE, standard hydrogen electrode; SWCNT,
single walled carbon nanotubes; TCA, trichloroacetic acid; TCEP,
Tris(2-carboxyethyl)phosphine.
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Stopped flow spectrophotometry. A DX.17MV sequential stopped-
flow spectrometer (Applied Photophysics, Leatherhead, U.K) with
a 3 ms deadtime was used to measure the single turnover oxidation
experiments of MsCDH by PMFT and DCPIP. All reactions were
carried out in 50 mM Hepes, pH 7.0, 250 mM NaCl, and 10% glyc-
erol. For reactions with PMFT, the absorbance of the MsCDH
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bound NADH was monitored at 340 nm (DeNADH 340nm = 6.22 mM-
1cm-1). For reactions with DCPIP, the absorbance of the oxidant was
monitored at 600 nm ( DeDCPIP 600nm = 20.7 mM-1cm-1). Reactions
were set up so that the final concentrations were 160 µM MsCDH
and 25 or 50 µM PMFT, or 22 or 60 µM DCPIP. Reactions were
run in triplicate at each concentration, averaged, and fitted to a single
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Supporting Information
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Corresponding Author
* J. A. Latham. E-mail: john.latham@du.edu
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Bacteriocin with a Narrow Spectrum of Activity against Clostridium Difficile.
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John A. Latham: 0000-0002-3034-1768
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Shanahan, F.; Kiely, B.; Hill, C.; Ross, R. P. Effect of Broad- and Narrow-
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Funding Sources
This work was supported by National Institutes of Health Grant GM
124002 to J.A.L.
ACKNOWLEDGMENT
We thank Dr. Monika Dzieciatkowska (University of Colorado Anchutz
Medical Campus) for performing the HRMS measurements.
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ABBREVIATIONS
AHDP, 3-amino-5-[(p-hydroxyphenyl) methyl]-4,4-dimethyl-2-
pyrrolidinone;
CHAPS,
(3-[(3-cholamidopropyl)dime-
thylammonio]-1-propanesulfonate; COSY, correlation spectros-
copy; CV, cyclic voltammetry; DCPIP, 2,3-dichlorophenolindophe-
nol; DTT, dithiothreitol; FAD, flavin adenine dinucleotide; FMN,
flavin mononucleotide; FPLC, fast protein liquid chromatography;
HEPES, 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid;
HMBC, heteronuclear multiple bond correlation; HPLC, high per-
formance liquid chromatography; HRMS, high resolution mass
spectrometry; HSQC, heteronuclear single quantum coherence;
IPTG, isopropyl-b-D-1-thiogalctopyanoside; MFT, mycofactocin;
MOPS, (3-(N-morpholino)propanesulfonic acid); NAD+/NADH,
nicotinamide dinucleotide; NMR nuclear magnetic resonance; PBS,
System?
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TOC FIGURE
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102
4
0
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-5 102
-0.50 -0.25 0.00
0.25
0.50
AHDP
Potential (V) vs SHE
+ MsMftD
8
+ MuMftD
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Retention Time (min)
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