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Physiology, Genomics, and Pathway Engineering of an Ethanol-
Tolerant Strain of Clostridium phytofermentans
Andrew C. Tolonen,
a
Trevor R. Zuroff,
b
*Mohandass Ramya,
c
Magali Boutard,
a
Tristan Cerisy,
a
Wayne R. Curtis
b
Genoscope-CEA, CNRS-UMR8030, Université d’Évry, Évry, France
a
; Department of Chemical Engineering, The Pennsylvania State University, University Park, Pennsylvania,
USA
b
; Department of Genetic Engineering, SRM University, Kattankulathur, India
c
Novel processing strategies for hydrolysis and fermentation of lignocellulosic biomass in a single reactor offer large potential
cost savings for production of biocommodities and biofuels. One critical challenge is retaining high enzyme production in the
presence of elevated product titers. Toward this goal, the cellulolytic, ethanol-producing bacterium Clostridium phytofermen-
tans was adapted to increased ethanol concentrations. The resulting ethanol-tolerant (ET) strain has nearly doubled ethanol tol-
erance relative to the wild-type level but also reduced ethanol yield and growth at low ethanol concentrations. The genome of the
ET strain has coding changes in proteins involved in membrane biosynthesis, the Rnf complex, cation homeostasis, gene regula-
tion, and ethanol production. In particular, purification of the mutant bifunctional acetaldehyde coenzyme A (CoA)/alcohol
dehydrogenase showed that a G609D variant abolished its activities, including ethanol formation. Heterologous expression of
Zymomonas mobilis pyruvate decarboxylase and alcohol dehydrogenase in the ET strain increased cellulose consumption and
restored ethanol production, demonstrating how metabolic engineering can be used to overcome disadvantageous mutations
incurred during adaptation to ethanol. We discuss how genetic changes in the ET strain reveal novel potential strategies for im-
proving microbial solvent tolerance.
The conversion of lignocellulosic biomass to fuels and com-
modities represents a large-scale, renewable alternative to pe-
troleum. This multistep bioconversion is traditionally performed
in a series of independent processes, but consolidated bioprocess-
ing (CBP) is an alternative paradigm with potential economic ad-
vantages (1). In CBP, enzyme production, hydrolysis, and fer-
mentation occur in a single reactor, leading to savings in capital
and operating costs as well as increased efficiencies due to system
synergies (2). Here we studied Clostridium phytofermentans,a
promising CBP candidate that ferments plant biomass primarily
to ethanol (3,4). C. phytofermentans hydrolyzes pretreated corn
stover (both glucans and xylans) with efficiencies similar to those
seen with simultaneous saccharification and cofermentation
(SSCF) using commercial enzymes and xylose-fermenting yeast
(Saccharomyces cerevisiae)(
5). Fermentation of pretreated corn
stover by C. phytofermentans reaches a titer of 7 g liter
⫺1
ethanol
(6), and stable cocultures of C. phytofermentans and S. cerevisiae
cdt-1 ferment ⬃70 g liter
⫺1
cellulose to 22 g liter
⫺1
ethanol (7),
which is an ethanol concentration that reduces C. phytofermentans
growth. Thus, application of CBP bacteria such as C. phytofermen-
tans will likely require improving their solvent tolerances without
compromising enzyme production or fermentation of soluble
carbohydrates to ethanol.
Considerable effort has focused on adapting clostridia to
increased solvent levels and investigating the genetic and phys-
iological changes associated with adaptation to solvents (8–
13). Other studies have shown increased ethanol production in
clostridia that primarily produce fermentation products other
than ethanol. C. cellulolyticum expressing pyruvate decarboxylase
and alcohol dehydrogenase (ADH) overcame pyruvate accumu-
lation and shifted fermentation products from lactate to acetate
and ethanol (14). In C. thermocellum, redirection of carbon flow
through pyruvate kinase (15), inactivation of lactate dehydroge-
nase and phosphotransacetylase (16), and deletion of hydroge-
nases (17) all improve ethanol production. These results demon-
strate that, although the genetic tools are being developed only
now, engineering improved ethanol production in cellulolytic
clostridia is possible. However, development of strains that are
ethanol tolerant (ET) and that also produce ethanol in high titers
remains a significant challenge.
Here we sought to develop a strain of C. phytofermentans with
both improved resistance and production of ethanol, particularly
from cellulose. We isolated an ethanol-tolerant (ET) C. phytofer-
mentans strain by serial transfer into increasing ethanol levels and
characterized its growth and fermentation properties. We se-
quenced the ET strain genome to reveal genomic mutations that
arose during adaptation and overcame reduced ethanol yield in
the ET strain by heterologous expression of an alternative ethanol
formation pathway. We discuss how the findings from this study
improve our understanding of how microbes adapt to elevated
concentrations of solvents such as ethanol.
Received 23 February 2015 Accepted 27 May 2015
Accepted manuscript posted online 5 June 2015
Citation Tolonen AC, Zuroff TR, Ramya M, Boutard M, Cerisy T, Curtis WR. 2015.
Physiology, genomics, and pathway engineering of an ethanol-tolerant strain of
Clostridium phytofermentans. Appl Environ Microbiol 81:5440 –5448.
doi:10.1128/AEM.00619-15.
Editor: J. L. Schottel
Address correspondence to Andrew C. Tolonen, atolonen@genoscope.cns.fr.
*Present address: Trevor R. Zuroff, Biodomain, Shell Technology Center, Houston,
Texas, USA.
A.C.T. and T.R.Z. contributed equally to this article.
Supplemental material for this article may be found at http://dx.doi.org/10.1128
/AEM.00619-15.
Copyright © 2015, American Society for Microbiology. All Rights Reserved.
doi:10.1128/AEM.00619-15
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MATERIALS AND METHODS
Culturing. C. phytofermentans ISDg (ATCC 700394) was grown anaero-
bically by preparing cultures in a Coy anaerobic chamber with a 1.5%
H
2
/98.5% N
2
atmosphere. Cultures were incubated without shaking at
30°C in GS2 medium (18) adjusted to a pH of 7 and supplemented with
carbon sources as described elsewhere in the text. Growth kinetics were
monitored by optical density at 600 nm (OD
600
) in sealed 100-well mi-
crotiter plates (Bioscreen 9502550) as previously described (19); cultures
were briefly shaken to resuspend cells before each optical density mea-
surement. Cellulose, cellobiose, and glucose cultures for substrate con-
sumption and fermentation product analysis were grown in 100-ml se-
rum bottles, which were sealed with butyl rubber stoppers after degassing.
The C. phytofermentans ethanol-tolerant (ET) strain was selected by
serial transfer (1:50 dilution) into culture tubes containing 10 ml of GS2
medium supplemented with increasing ethanol concentrations. Starting
with cultures in 4% (vol/vol) (31.5 g liter
⫺1
) ethanol, cultures were trans-
ferred weekly to fresh medium containing the same ethanol concentration
and to medium with a 1%-higher ethanol concentration. If no growth was
observed at the higher concentration after 1 week, cultures were retrans-
ferred to the same ethanol concentrations. If a culture grew at a higher
ethanol concentration, this culture was transferred again to that ethanol
concentration and to a 1%-greater ethanol concentration. Growth was
observed at 5% ethanol after 7 weekly transfers, 6% after 13 transfers, and
7% after 19 transfers. Each time the ethanol tolerance improved, cells were
TABLE 1 Bacterial strains, primers, and plasmids used in this study
Strain name Genotype or description
a
Source or sequence
Strains
Clostridium
phytofermentans ISDg
ATCC type strain 700394 Susan Leschine Laboratory, University of Massachusetts, Amherst, Amherst,
MA, USA
E. coli Top 10 hsdR mcrA endA1 recA1 rpsL (Str
r
)
(cloning strain)
Invitrogen Corporation
E. coli S17-1 RP4-2 (Km::Tn7Tc::Mu-1) recA1
endA1 (conjugal strain)
Yale E. coli Stock Center
Primers
pdcAdhB_F Forward primer for amplification of pdc
and adhB from pES120
5=-TTTTTCGAATTCACCGGATCCCTGCAGTAGGAGGAATTAACC-3=
pdcAdhB_R Reverse primer for amplification of pdc
and adhB from pES120
5=-ATATTTCGATCGATTGCATGCTTAGAAAGCGCTCAGGAAGAG-3=
pQexp_F Forward primer to confirm pdc-adhB
insertion in pQexp
5=-AAACCTAGGTAATTGAGGAAAGTTACAATTA-3=
pQexp_R Reverse primer to confirm pdc-adhB
insertion in pQexp
5=-GAATGGCGCCTGATGCG-3=
cphy3925F Forward primer to amplify Cphy3925
coding sequence
5=-AAAGAAGGAGATAGGATCATGACGAAGAAAGTGGAATTA-3=
cphy3925R Reverse primer to amplify Cphy3925
coding sequence
5=-GTGTAATGGATAGTGATCTTAATGGTGATGGTGATGATGTTTACCG
TAGTACACTTTAAGATAG-3=
Plasmids
pES120 Source of Z. mobilis pdc and adhB genes Jay Keasling Laboratory, University of California, Berkeley, Berkeley, CA, USA
pQexp Replicating plasmid for C.
phytofermentans
Andrew Tolonen Laboratory, Genoscope-CEA, Évry, France
pQexpE pQexp with Z. mobilis pdc and adhB
cloned into the unique BamHI and
PvuI sites
This study
a
Str, streptomycin, Km, kanamycin; Tc, tetracycline.
FIG 1 Growth of wild-type (WT) (A) and ethanol-tolerant (ET) (B) C. phytofermentans strains at 30°C in GS2 medium with 3 g liter
⫺1
glucose supplemented
with ethanol at the following levels (vol/vol): 0% (red squares), 2% (green circles), 4% (blue triangles), 6% (yellow diamonds), and 7% (black X’s). Data points
of growth (OD
600
) represent means of results from triplicate 400-l cultures in sealed microtiter plates; error bars represent 1 standard deviation.
C. phytofermentans Ethanol Tolerance and Production
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plated, individual colonies were picked, and liquid cultures were reinoc-
ulated to ensure that selection was based on a specific strain with increased
ethanol tolerance and not on a consortium of strains that collectively
survived the increased ethanol concentration. The ET strain is thus a
colony-purified isolate from a mother culture that grew in GS2 medium
supplemented with 7% (vol/vol) ethanol. After colony purification, it was
confirmed the ET strain has ethanol resistance similar to that of the
mother culture.
Cellulose and fermentation analysis. The level of cellulose remaining
in the culture was measured by taking a 1-ml sample from a 10-ml culture
tube with a sterile syringe, placing it in a preweighed 1.7-ml microcentri-
fuge tube, and centrifuging at 13,000 ⫻gfor 10 min. The supernatant was
removed, and the cellulose pellet was washed and centrifuged again at
13,000 ⫻gfor 10 min. The rinsed pellet was placed at 70°C to dry until a
constant mass was reached. The contribution of cellular biomass to total
cellulose weight was not accounted for and was assumed to be minimal
due to low anaerobic biomass yields.
Fermentation product concentrations were measured in 0.22-m-pore-
size-filtered culture supernatant using an Agilent 1100 high-performance liq-
uid chromatograph (HPLC) with a Jasco RI-1531 refractive index detector
(RID) and an Aminex HPX-87H cation exchange column (Bio-Rad). The
HPLC was run using a 0.01 M sulfuric acid mobile phase, 65°C column tem-
perature, 30°C RID temperature, 25 l sample volume, and 0.6 ml/min op-
erating flow rate. Product formation is reported relative to the concentration
in the medium at the point of inoculation. Gas phase measurements were
made by removing 1 ml of headspace and injecting 100 l into a gas chro-
matograph (Model 8610C multiple-gas analyzer; SRI Instruments). Argon
was used as a carrier gas and was adjusted to 30 lb/in
2
gauge pressure. A
stainless steel molecular sieve (13⫻) and silica gel-packed columns were used
for sample separation, and the components were detected using a thermal
conductivity detector (TCD). The column compartment temperature was
held initially at 40°C for 3.5 min and then ramped to 160°C for 2 min and to
300°C for 10 min, after which the column was allowed to cool to 40°C for the
remainder of the sample run.
Genome sequencing and variant analysis of the ET strain. A total of
12 g of genomic DNA was extracted from a 4-ml ET strain culture using
a Sigma GenElute bacterial genomic DNA kit (NA2110). DNA was se-
quenced on an Illumina MiSeq instrument with an insertion size of 795 bp
FIG 2 C. phytofermentans yields (mole of product per mole of glucose equivalent consumed) of ethanol from WT (dark red) and ET (light red) strains (A) and
acetate from WT (dark blue) and ET (light blue) strains (B). Statistical differences between WT and ET yield averages are indicated at Pvalues of ⬍0.01 (*), ⬍0.05
(**), and ⬍0.1 (***) using Student’s ttest. (C) C. phytofermentans WT and ET cellobiose consumption and fermentation products in GS2 cellobiose medium with
0, 3.75, or 7% (vol/vol) added ethanol. Bars show concentrations (in millimoles) of cellobiose consumption (gray) and production of ethanol (red), acetate
(blue), and formate (yellow) by WT (dark bars) and ET (light bars) strains. All cultures were grown in serum bottles containing GS2 medium at 30°C with 30g
liter
⫺1
cellulose, 10 g liter
⫺1
cellobiose, or 30 g liter
⫺1
glucose and measured after 14 days. Bars represent averages of the results from 4 cultures; error bars
represent 1 standard deviation.
Tolonen et al.
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and 300-bp paired-end reads. A total of 5,077,282 reads passed quality
filtering and mapping by Picard Tools (https://github.com/broadinstitute
/picard), yielding approximately 250-fold genome coverage. Sequence
variants (single nucleotide polymorphisms [SNPs] and indels) in the ET
genome relative to the reference strain genome (NCBI accession no.
NC_010001.1) were identified using the Genome Analysis Toolkit
(GATK) (20) (see the supplemental material for detailed descriptions of
the filtering and variant-calling methods).
ADH purification and activity measurements. The Cphy3925-en-
coding genes from the wild-type (WT) and ET strains were cloned by
ligation-independent cloning (21) into pET-22B(⫹)as previously de-
scribed (19). Genes were cloned with C-terminal His tags using primer
pair cphy3925F/cphy3925R (Table 1) and confirmed by sequencing. Plas-
mids were transformed into Escherichia coli BL21(DE3) (Novagen 70235)
and grown in 50 ml TB medium (12 g liter
⫺1
tryptone, 24 g liter
⫺1
yeast
extract, 4 ml liter
⫺1
glycerol) to an OD
600
of 1, and expression was in-
duced by adding 500 M IPTG (isopropyl--D-thiogalactopyranoside)
and incubating overnight at 20°C. Cells were pelleted, resuspended in lysis
buffer (50 mM phosphate buffer [pH 8], 0.5 M NaCl, 10 mM imidazole,
15% glycerol, 1 mM Pefabloc [Sigma 76307]), and lysed by sonication
(Cole-Parmer Vibracell CV33) with lysozyme (Novagen 71230). His-
tagged proteins were purified from 50 ml culture on nickel-nitrilotriacetic
acid (Ni-NTA) spin columns (Qiagen 31014) and visualized on 12% SDS-
PAGE gels (Novex 12% Bis-Tris gel NP0342BOX). Enzyme activities were
measured as described in reference 22 in 100 lof 100 mM Tris-HCl (pH
8) containing cofactor [0.2 mM NADH(P)H or 2 mM NAD(P)
⫹
] and
substrate (300 M acetyl coenzyme A [acetyl-CoA] or CoA, 18 mM acet-
aldehyde, 2 M ethanol) as appropriate. NAD(P)H was measured as
340-nm absorbance (extinction coefficient, 6.22 mM
⫺1
·cm
⫺1
) using a
SAFAS UVmc2 spectrophotometer at room temperature.
Plasmid construction and transformations. pQexpE is derived from
pQexp (23), a plasmid that replicates stably with erythromycin selection
in E. coli (200 gml
⫺1
erythromycin) and C. phytofermentans (40 g
ml
⫺1
erythromycin on plates and 200 gml
⫺1
erythromycin in liquid
culture). To construct pQexpE, the Z. mobilis pdc and adhB genes were
PCR amplified from pES120 (24) using primer pair pdcAdhB_F/dcAd-
hB_R (Table 1) and cloned into the unique BamHI and PvuI sites of
pQexp. The insertion was confirmed by sequencing using primer pair
pQexp_F/pQexp_R. pQexpE was conjugally transferred to C. phytofer-
mentans using donor strain E. coli S17-1. Conjugation was performed as
described in references 23 and 25, except polyethersulfone membranes
were used to support 50-l culture mixtures and only nalidixic acid with-
out trimethoprim was used to select against E. coli following mating. Pos-
itive C. phytofermentans transconjugants containing pQexpE were con-
firmed by colony PCR using primer pair pQexp_F/pQexp_R to amplify
the 2.9-kb pdc-adhB operon.
Nucleotide sequence accession number. The FASTQ-formatted
DNA sequencing files for the ET genome were submitted to the European
Nucleotide Archive under primary accession no. PRJEB7255.
RESULTS AND DISCUSSION
Isolation and physiology of an ethanol-tolerant C. phytofer-
mentans strain. The growth of wild-type (WT) C. phytofermen-
TABLE 2 Genomic DNA variants in the ET strain
a
Protein
Length
(amino acids) Amino acid variant
Confidence (Q)
value Annotation
Energy and metabolism
Cphy3925 (AdhE) 872 G609D 8,929 Fe-dependent bifunctional acetaldehyde-CoA/alcohol dehydrogenase
Cphy0215 (RnfA) 191 C26S 5,569 Ferredoxin:NAD
⫹
oxidoreductase (Fno); couples electron transfer
from reduced ferredoxin to NAD
⫹
with cation transport out of the
cell to create an electrochemical gradient
Cphy3255 259 20-bp insertion at
Q91
15,938 FMN-dependent nitroreductase
Transport
Cphy0543 (MgtA/MgtB) 920 K417N 8,661 P-type ATPase for Mg
2⫹
uptake transporter or Ca
2⫹
/Mg
2⫹
antiporter
Cphy3778 258 1-bp deletion, Y239
frameshift
11,687 Na
⫹
efflux transporter, ABC-2 transmembrane permease (PFAM
accession no. PF06182)
Membrane and cell wall
Cphy0233 (PlsD) 237 D80N 9,246 Membrane synthesis: glycerol-3-phosphate acyltransferase; transfers
a fatty acid to the 1 position of glycerol-3-phosphate
Cphy0107 (MurC) 469 G115D 8,841 Peptidoglycan synthesis: ATP-dependent ligation of L-alanine and
UDP-N-acetylmuramic acid to form
UDP-N-acetylmuramyl-L-alanine
Gene regulation
Cphy3040 301 D182N 8,214 LysR transcriptional activator/repressor
Cphy3687 (PolB) 1,278 D189Y 8,824 -Subunit of DNA-directed RNA polymerase
Noncoding changes
Cphy0267 524 Q65Q (synonymous) 7,470 Modification methyltransferase
Intergenic G¡T
transversion
None 52 Transversion between 2 convergently transcribed genes, encoding
Cphy1313 and Cphy1314
Cphy3036 (ApbE) 381 L244L (synonymous) 9,011 Thiamine biosynthesis: membrane-associated lipoprotein
a
Data include the gene name, encoded protein length, amino acid (if coding) or DNA variant, confidence value (Q) of the variant call, and annotation of the mutated protein. The
probability (P) that a variant exists in the genome is reported as a Phred-scaled probability, i.e., Q⫽⫺10 ⫻log
10
(1 ⫺P), meaning that a Qvalue of 100 indicates an error
probability (1 ⫺P)of10
⫺10
(see GATK reports in the supplemental material for more information). FMN, flavin mononucleotide.
C. phytofermentans Ethanol Tolerance and Production
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tans was monitored in cultures supplemented with 0, 2, 4, 6, or 7%
(vol/vol) ethanol (Fig. 1A), and generation times (hours) and
maximum cell densities (OD
600
) were calculated (see Table S1 in
the supplemental material). The WT strain cultures grew similarly
at 0% ethanol and 2% ethanol but growth was significantly inhib-
ited at 4% ethanol, and very little growth was observed at 6 and 7%
ethanol. In contrast, the ET strain grew at 4, 6, and 7% ethanol,
with a maximum cell density (OD
600
) at 7% ethanol similar to that
of the WT strain at 4% ethanol (Fig. 1B; see also Table S1).
While the ET strain was more ethanol resistant, it grew more
slowly than the WT strain and grew at reduced cell yields in cul-
tures without added ethanol (Fig. 1). Glucose consumption by the
ET strain was also slower than glucose consumption by the WT
strain, reflecting the reduced growth rate. When grown on 30 g
liter
⫺1
glucose, the ET strain consumed only 20 g liter
⫺1
substrate
in 200 h, while the WT strain completely exhausted the glucose in
less than 150 h (see Fig. S1 in the supplemental material). Thus,
the enhanced growth of the ET strain at a high ethanol concentra-
tion is accompanied by reduced growth under standard condi-
tions, supporting the idea that the ET strain has altered physiol-
ogy, with results similar to those seen with C. thermocellum
adapted to 5% ethanol (26,27). However, we observed no mor-
phological differences between ET and WT colonies grown on
GS2 agar plates or cells grown in liquid GS2 medium (see Fig. S2).
The ET strain also produces less ethanol per unit of sugar con-
sumption than the WT strain. For example, ethanol yield (moles
of product per mole of glucose equivalent consumed) by the ET
strain decreased 25 to 50% relative to that seen with the WT strain
when growing on cellulose, cellobiose, and glucose (Fig. 2A),
whereas the acetate yields of the strains were similar (Fig. 2B). We
also measured cellobiose consumption and formation of the pri-
mary fermentation products ethanol, acetate, and formate in WT
and ET cultures grown in cellobiose medium supplemented with
0, 3.75, or 7% ethanol (Fig. 2C). Ethanol production by the WT
strain was significantly lower at 3.75% ethanol, and cellobiose
consumption and fermentation ceased at 7% ethanol. In contrast,
cellobiose consumption and ethanol production by the ET strain
decreased only slightly with increased ethanol supplementation,
demonstrating the robustness of the ethanol resistance pheno-
type. In all, these results highlight the phenotypic advantages of
the ET strain with respect to metabolizing and producing ethanol
at elevated ethanol concentrations but also indicate that those
advantages occur at the expense of lower ethanol yields and slower
growth at low ambient concentrations of ethanol.
Genome sequence of the ET strain. The C. phytofermentans ET
strain genome contains 12 variants relative to the WT strain (Ta-
ble 2), many fewer than a C. thermocellum ET genome with similar
ethanol resistance that had 200 to 500 changes (9). While ethanol
resistance is likely a complex, multigenic trait, the small number of
changes in the ET strain genome shed light on DNA variants that
could have functional roles in ethanol tolerance. Two of the 12
mutations were in genes encoding transcriptional regulators that
could effectuate broad gene expression changes: PolB, the -sub-
unit of DNA-directed RNA polymerase, and a LysR regulator,
Cphy3040. LysR-type regulators often colocalize in the genome
with their targets (28) and the gene encoding Cphy3040 is adja-
cent to a gene encoding a NAD-dependent aldehyde dehydroge-
nase, suggesting that this regulator is related to alcohol formation.
Ethanol increases the permeability of the cell membrane, re-
sulting in toxic leakage of metabolites out of the cell (29). Ethanol
resistance thus often involves membrane modifications such as
altered protein content (8) or longer chain fatty acids and more
plasmalogen lipids (30) that increase membrane rigidity to miti-
gate the fluidizing effect of ethanol. The ET strain has a D80N
change in the putative acyl-acceptor binding pocket (NCBI acces-
sion no. cd07989) of Cphy0233, a homolog of C. butyricum PlsD
that transfers a fatty acyl group to the sn-1 position of glycerol-3-
phosphate in phospholipid biosynthesis (31). The D80N
FIG 3 Comparison of activities of purified Cphy3925 AdhE from wild-type and ET strains. (A) SDS-PAGE gel of purified Cphy3925 from the wild-type (WT)
and ET (G609D) strains showing single bands of the expected 95-kDa molecular mass. (B to E) Reactions for the two-step, bidirectional interconversionof
acetyl-CoA acetaldehyde and ethanol: reduction of 300 M acetyl-CoA to acetaldehyde (red) (B), reduction of 18 mM acetaldehyde to ethanol (green) (C),
oxidation of 2 M ethanol to acetaldehyde (purple) (D), and oxidation of 18 mM acetaldehyde to acetyl-CoA (blue) (E). Enzyme activities are shown in millimoles
of NAD(P)H per micromole of enzyme per second measured using NADH(P)H or NAD(P)
⫹
cofactors. Bar heights represent averages of duplicate activity
measurements, and error bars represent 1 standard deviation.
Tolonen et al.
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Cphy0233 mutation may thus enable synthesis of a more rigid,
ethanol-resistant cell membrane by altering which fatty acids are
incorporated into phospholipids.
The ET strain has a C26S mutation in the RnfA subunit of the
membrane-bound Rnf complex that couples efflux of H
⫹
(32)or
Na
⫹
(33) with electron transfer from reduced ferredoxin to
NAD
⫹
(34). The resulting electrochemical gradient is harnessed
by an F
0
F
1
ATPase for ATP synthesis. The C. phytofermentans Rnf
complex (Cphy0211 to Cphy0216) and F
o
F
1
ATPase (Cphy3735
to Cphy3742) are highly expressed on all tested carbon sources
(19) and may be important for energy conservation, similarly to C.
ljungdahlii (32). However, Rnf generates NADH, which may not
be tolerated by the ET strain that cannot reoxidize NADH by
AdhE-mediated ethanol formation (see below). The C26S RnfA
variant may thus cripple the Rnf complex, which sacrifices ATP
production, but may benefit the ET strain by balancing cellular
NADH/NAD
⫹
ratios.
The ET strain also has mutations in two transporters putatively
involved in cation homeostasis. Cphy0543 is homologous to
MgtA, a P-type ATPase upregulated at low ambient Mg
2⫹
concen-
trations (35) to mediate Mg
2⫹
uptake (36)orCa
2⫹
/Mg
2⫹
antiport
(37). Cphy3778, the membrane component of an ABC trans-
porter (PFAM accession no. PF06182), appears to be cotrans-
cribed with Cphy3780, an ABC-type Na
⫹
efflux protein (NCBI
accession no. cd03267). In Bacillus subtilis, this Na
⫹
efflux system
is induced by ethanol and is proposed to compensate for an influx
of extracellular Na
⫹
resulting from a weakened membrane barrier
(38). The variants in these cation transporters may increase their
activities to alleviate cation leakage due to ethanol stress.
AdhE activities. The ET strain has a G609D variant in
Cphy3925 AdhE, a putative acetaldehyde-CoA dehydrogenase
and alcohol dehydrogenase (ADH). The G609D mutation is in a
conserved position in the active site of the C-terminal ADH do-
main (NCBI accession no. cd08178). A previous study reported an
ethanol-tolerant C. thermocellum strain with AdhE mutations
(P704L and H735R) that shifted the cofactor specificity from
FIG 4 (A) Diagram of C. phytofermentans carbon metabolism showing insertion of the Z. mobilis pdc-adhB alternative ethanol formation pathway. Enzymatic
steps: {a}, lactate dehydrogenase (Cphy1117); {b}, pyruvate formate lyase (Cphy1174); {c}, pyruvate ferredoxin oxidoreductase (Cphy3558); {d}, Rnf ferredoxin:
NAD
⫹
oxidoreductase complex (Cphy0211 to Cphy0216); {e}, phosphate acetyltransferase (Cphy1326); {f}, acetate kinase (Cphy1327); {g}, acetaldehyde
dehydrogenase (Cphy1428 or Cphy3925); {h}, alcohol dehydrogenase (Cphy3925 or Cphy1029); {i}, Zymomonas mobilis pyruvate decarboxylase (pdc); {j},
alcohol dehydrogenase (adhB). Dashed lines represent multienzyme reactions where all enzymes are not listed. (B) Plasmid map of pQexpE for pdc-adhB
expression in C. phytofermentans. Plasmid features: Gram-negative pUC origin of replication (oriR pUC), C. phytofermentans pyruvate ferredoxin oxidoreduc-
tase promoter (P3558) to express the Z. mobilis pdc-adhB genes, the RP4 conjugal origin of transfer (oriT), the Gram-negative/Gram-positive erythromycin
resistance gene from TN1545 (ermR), the Gram-positive pAMB1 origin (oriR pAMB1), and repE-encoded protein.
FIG 5 Cellulose consumption (A) and ethanol production (B) by non-plas-
mid-bearing (⫺) and pdc-adhB-containing (⫹) WT and ET cultures at 30°C in
GS2 medium with 3 g liter
⫺1
␣-cellulose after 30 days. Bars represent averages
of the results from duplicate cultures in serum bottles from three independent
experiments, and error bars represent 1 standard deviation. Statistical differ-
ences between treatments are indicated at Pvalues of ⬍0.01 (*) and ⬍0.05 (**)
using Student’s ttest.
C. phytofermentans Ethanol Tolerance and Production
August 2015 Volume 81 Number 16 aem.asm.org 5445Applied and Environmental Microbiology
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NADH to NADPH, which was proposed to confer ethanol re-
sistance by altering the internal redox balance (9). To deter-
mine the effect of the G609D mutation on Cphy3925 enzyme
activity, we purified WT and ET versions of the enzyme (Fig.
3A) and tested their in vitro catalysis of the two-step, bidirec-
tional reactions converting acetyl-CoA to ethanol using either
NADH or NADPH cofactors.
The mutated Cphy3925 lost NAD(H)-dependent activities,
but, unlike the mutated AdhE in C. thermocellum, the G609D
mutation did not result in NADPH-dependent ADH activity (Fig.
3B to E). Instead, our results support the notion that the ET strain
arrested AdhE-mediated interconversion of acetyl-CoA, acetalde-
hyde, and ethanol, which helps explain why the C. phytofermen-
tans ET strain had lower ethanol yield. AdhE loss of function could
mitigate ethanol stress by reducing intracellular levels of ethanol
and its highly toxic precursor, acetaldehyde. C. phytofermentans
encodes four Fe-dependent ADHs in addition to Cphy3925 as well
as a Zn-dependent ADH. All 6 ADHs are expressed, and
Cphy3925 and Cphy1029 are among the most highly expressed
proteins on all tested carbon sources (19,39). C. phytofermentans
thus likely produces ethanol by the concerted action of multiple
ADHs, and these other ADHs, especially Cphy1029, are responsi-
ble for ethanol produced by the ET strain.
Ethanol pathway engineering. To augment ethanol produc-
tion by the ET strain, an alternative ethanol production path-
way comprised of pyruvate decarboxylase (Pdc) and alcohol
dehydrogenase (AdhB) from Zymomonas mobilis (Fig. 4A) was
transferred into C. phytofermentans on the replicating pQexpE
plasmid (Fig. 4B). Together, these enzymes couple decarboxyl-
ation of pyruvate to ethanol with the oxidation of NADH and thus
represent an alternative to the AdhE ethanol formation pathway.
We chose to express foreign enzymes rather than a WT copy of
Cphy3925 because AdhE multimerizes (40) such that the mutant
AdhE could have a dominant-negative effect in a merodiploid.
Expression of pQexpE increased cellulolysis by ⬃30% in both
the WT and ET strains (Fig. 5A) and boosted ethanol production
by 70% relative to the ET strain (P⬍0.01), thereby restoring
ethanol yields to WT levels (Fig. 5B). CO
2
production increased
disproportionately relative to H
2
production in WT and ET
strains expressing pQexpE (Table 3). Elevated CO
2
synthesis is
likely due to increased pyruvate decarboxylation by the Pdc en-
zyme. Previous results showed that Pdc/AdhB expression en-
hanced cellulolysis and ethanol production in WT C. cellulolyti-
cum, which was proposed to result from consumption of excess
pyruvate that otherwise leads to metabolic arrest (14). Increased
metabolism (cellulolysis and production of CO
2
and ethanol) by
C. phytofermentans expressing Pdc/AdhB might be due to allevia-
tion of inhibition by excess pyruvate. Alternatively, expression or
activity of glycolytic enzymes might be regulated by NADH levels
such that NADH reoxidation by Pdc/AdhB stimulates glycolysis,
which results in increased substrate utilization.
Conclusions. In this study, we investigated the genetic basis
and phenotypic consequences of microbial ethanol tolerance by
isolating, characterizing, and engineering an ethanol-resistant
(ET) strain of Clostridium phytofermentans. The ET strain grows at
higher ethanol concentrations than the wild-type strain (Fig. 1)
and continues to produce ethanol at a 7% ambient ethanol
concentration (Fig. 2C) but has impaired growth (Fig. 1) and
ethanol yield (Fig. 2A) relative to the wild type. The genome
sequence of the ET strain revealed 12 mutations in genes in-
volved in diverse aspects of metabolism (Table 2), including a
G609D variant in the bifunctional acetaldehyde CoA/alcohol
dehydrogenase AdhE that abolishes its activity (Fig. 3). We
complemented the AdhE mutation in the ET strain by express-
ing pyruvate decarboxylase (Pdc) and alcohol dehydrogenase B
(AdhB) from Zymomonas mobilis on the pQexpE plasmid (Fig.
4), which boosted substrate conversion (Fig. 5A) and restored
ethanol production (Fig. 5B).
Additional work is needed to enhance C. phytofermentans’
plant biomass degradation and ethanol formation rates and prod-
uct titers. Recently, improvement of C. phytofermentans’ growth
on cellobiose, cellulose, and xylan by experimental evolution
yielded strains that also produced ethanol more quickly (41). The
genome sequence of the ET strain presented here suggests other
novel approaches to potentially improve ethanol resistance and
production. For example, our results suggest that further studies
on ethanol resistance should focus on PlsD-mediated fatty acid
incorporation into phospholipids, LysR-regulated gene expres-
sion patterns, overexpression of the Rnf complex to stimulate
AdhE-mediated ethanol production, and prevention of cation
leakage.
ACKNOWLEDGMENTS
C. phytofermentans ISDg was kindly provided by Susan Leschine (Univer-
sity of Massachusetts, Amherst).
A.C.T. was supported by a CNRS Chaire d’Excellence and the Geno-
scope-CEA. T.R.Z. was supported by a GRFP Fellowship from the Na-
tional Science foundation (grant no. DGE1255832) and a John and Jean-
nette McWhirter Fellowship from The Pennsylvania State University.
M.R. was supported by the Faculty Abroad Program, SRM University.
Anaerobic growth capabilities for this work were financed by the Geno-
scope-CEA and a grant from the Department of Energy Advanced Re-
search Project Agency—Energy (ARPA-e; no. DE-AR0000092) to W.R.C.
We thank Alex Rajangam for facilitating M.R.’s international ex-
change, Sergio Florez for useful feedback, Patrick Hillery and Sam
Bjork for experimental assistance, and Nymul Khan for supporting GC
analysis. We acknowledge the Huck Institute of the Life Sciences
Shared Fermentation Facility for providing HPLC analytical capabili-
ties and the Genoscope-CEA Sequencing Platform for sequencing the
ET strain. Finally, we thank George Church and Noel Goddard for
facilitating a sabbatical leave for W.R.C. at Harvard, where these stud-
ies were initiated.
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