Engineering the bioconversion of methane andmethanol to fuels and chemicals in native and syntheticmethylotrophsR Kyle Bennett1,2, Lisa M Steinberg1,2, Wilfred Chen1 andEleftherios T Papoutsakis1,2
Available online at www.sciencedirect.com
ScienceDirect
Methylotrophy describes the ability of organisms to utilize
reduced one-carbon compounds, notably methane and
methanol, as growth and energy sources. Abundant natural gas
supplies, composed primarily of methane, have prompted
interest in using these compounds, which are more reduced
than sugars, as substrates to improve product titers and yields
of bioprocesses. Engineering native methylotophs or
developing synthetic methylotrophs are emerging fields to
convert methane and methanol into fuels and chemicals under
aerobic and anaerobic conditions. This review discusses
recent progress made toward engineering native
methanotrophs for aerobic and anaerobic methane utilization
and synthetic methylotrophs for methanol utilization. Finally,
strategies to overcome the limitations involved with synthetic
methanol utilization, notably methanol dehydrogenase kinetics
and ribulose 5-phosphate regeneration, are discussed.
Addresses1Department of Chemical and Biomolecular Engineering, University of
Delaware, 150 Academy St., Newark, DE 19716, USA2The Delaware Biotechnology Institute, Molecular Biotechnology
Laboratory, University of Delaware, 15 Innovation Way, Newark, DE
19711, USA
Corresponding author: Papoutsakis, Eleftherios T ([email protected])
Current Opinion in Biotechnology 2018, 50:81–93
This review comes from a themed issue on Energy biotechnology
Edited by Akihiko Kondo and Hal Alper
https://doi.org/10.1016/j.copbio.2017.11.010
0958-1669/ã 2017 Elsevier Ltd. All rights reserved.
IntroductionAbundant natural gas supplies have made methane and
methanol promising substrates for biological production
of fuels and chemicals [1��]. These one-carbon (C1)
compounds are at least 50% more reduced than traditional
sugars, for example, glucose, allowing for improved prod-
uct titers and yields [2��]. Worldwide, the amount of
recoverable natural gas is estimated to be 7.2 � 103 tril-
lion ft3 [1��]. In the US alone, estimates approach
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2 � 103 trillion ft3. At current energy usage rates, this is
enough natural gas to supply the US for 100 years. Meth-
ane is also a potent greenhouse gas, having a warming
potential 21 times that of CO2. As a result, along with the
food versus fuel debate, biological gas-to-liquid (GTL)
conversion technologies are promising alternatives for
fuel and chemical production. This review discusses
recent progress made toward understanding and
engineering native methanotrophs and synthetic methy-
lotrophs for production of fuels and chemicals. Advance-
ments in aerobic and anaerobic methane utilization will
first be discussed, followed by those made toward engi-
neering synthetic methylotrophs for methanol utilization.
Finally, difficulties with engineering synthetic methanol
utilization and strategies to overcome them will be
detailed.
Aerobic methane utilization to produce fuelsand chemicalsThe physiology and biochemistry of aerobic methano-
trophs, which utilize methane as their sole carbon and
energy source, have been extensively reviewed [3,4]. The
first step in methane assimilation is oxidation to methanol
by methane monooxygenase (MMO) [5], followed by
oxidation to formaldehyde by pyrroloquinoline quinone
(PQQ)-containing methanol dehydrogenase (MDH)
[3,4]. Type I methanotrophs are gammaproteobacteria,
which assimilate formaldehyde via the ribulose monopho-
sphate (RuMP) pathway. Type II methanotrophs, which
assimilate formaldehyde via the serine cycle, are alpha-
proteobacteria [4]. A third group of aerobic methanotrophs,
Type X, utilize the RuMP pathway for formaldehyde
assimilation but express low levels of serine cycle enzymes
and grow at higher temperatures [3].
Two types of MMOs have been identified [5,6]. Nearly
all methanotrophs express a membrane-bound particulate
MMO (pMMO), and a few also express a soluble MMO
(sMMO). pMMO is an integral membrane hydroxylase
with three subunits arranged as an a3b3g3 trimer, encoded
by the pmoCAB operon, and contains two Cu-containing
active sites in the N-termini and C-termini of pmoB[6]. sMMO contains three components: a hydroxylase,
encoded by mmoX, mmoY and mmoZ, a reductase,
encoded by mmoC, and a regulatory protein, encoded
by mmoB [5,6]. The hydroxylase is an a2b2g2 dimer with
a diiron active site in the alpha subunit [5]. pMMO has a
Current Opinion in Biotechnology 2018, 50:81–93
82 Energy biotechnology
narrow substrate specificity and oxidizes shorter alkanes
up to five carbons [6] whereas sMMO has a broader
substrate range that includes aromatic and heterocyclic
compounds [6].
In addition to oxygen, some methanotrophs use alternate
electron acceptors for methane activation. A methane-
oxidizing, nitrite-reducing enrichment culture from fresh-
water sediment was dominated by one bacterial species
[7], and metagenomic sequencing led to the construction
of the full draft genome of a proposed new species,
Methylomirabilis oxyfera [8], which possesses a pMMO
and an incomplete denitrification pathway. Methane is
oxidized with nitrite and a pathway was proposed in
which two molecules of NO could be used to produce
N2 and O2 for methane oxidation [8]. Methane oxidation
coupled to nitrate reduction was described for Methylomonasdenitrificans under hypoxia [9].
There is increased interest in engineering methanotrophs
for converting methane into fuels and chemicals. Improv-
ing methane oxidation, either by MMO overexpression or
enhanced activity via protein engineering, could increase
efficiency. However, MMO expression in heterologous
hosts has largely failed [10]. A number of genetic tools
have been developed for methanotrophs [10], including
conjugation for introducing genetic material from E. coli.Methylomicrobium buryatense 5G is emerging as a tractable
host for metabolic engineering with advances including
engineering of a strain capable of IncP-based vector repli-
cation for episomal gene expression [11], development of
selection/counter-selection markers for allelic exchange
[11] and transformation using electroporation [10].
Currently, Methylosinus trichosporium is the preferred spe-
cies for methanol production, which requires a co-sub-
strate and inhibition of MDH [12]. Another strategy for
methanol production is co-feeding methane and ammonia
to a nitrifying culture where the methanol produced by
action of ammonia monooxygenase cannot be used by the
nitrifiers [13]. A third strategy uses an engineered BM-3
cytochrome P450 monooxygenase from Bacillus megateriumfor methane oxidation [14].
One product from methane is polyhydroxybutyrate
(PHB), a biopolymer and plastic substitute [15,16].
Methanotrophs synthesize intracellular PHB as a source
of reducing equivalents for growth under nutrient-limit-
ing conditions albeit yields are modest and of low molec-
ular weight [15,16]. Methylobacterium organophilum CZ-2
was reported to accumulate up to 57% PHB under nitro-
gen limitation [17]. Another product are storage lipids in
the form of triacylglycerides (TAGs), which can be con-
verted to biodiesel [18]. TAG accumulation is promoted
under oxygen-limiting, nitrogen-limiting or phosphate-
limiting conditions [18]. In some methanotrophs, carbon
flux can be routed through the phosphoketolase pathway
Current Opinion in Biotechnology 2018, 50:81–93
into lactic and acetic acid with increased ATP and
decreased CO2 production [19]. Overexpression of phos-
phoketolase in M. buryatense led to a 2.6-fold improve-
ment in biomass and lipid yield from methane [20�].
Methylomonas sp. 16a is an interesting methanotroph due
to high-level production of C30 carotenoids [21], but the
production of larger carotenoids remains challenging due
to lack of genetic tools. Episomal gene expression for
synthesis of C40 carotenoids, astaxanthin and canthaxan-
thin, resulted in yields of 2.4 g gDW�1 [22]. Increased
yields were obtained by optimizing chromosomal inte-
gration location [23] and co-expression of bacterial
hemoglobins [24].
Efforts have also been made to engineer methanotrophs
for high-volume chemicals, for example, lactic and suc-
cinic acids. Overexpression of lactate dehydrogenase
(LDH) from Lactobacillus helveticus in M. buryatenseimproved lactate production by 70-fold over the wild-
type strain, resulting in 0.8 g/L [25��]. Expression of the
succinate synthesis pathway in M. capsulatus Bath
resulted in 70 mg/L [26�]. Trace-level production of
1,4-butanediol [27�] and isobutanol [28�] has also been
reported.
Although aerobic methane conversion to fuels and che-
micals has been demonstrated, only low yields were
achieved at small scale. During the oxidation of methane
to methanol via MMO, two electrons are required to
simultaneously reduce O2 to H2O. Recovery of these
electrons is achieved in the subsequent step of methanol
oxidation to formaldehyde. Therefore, the result is the
redox-neutral conversion of methane to formaldehyde,
which results in a 36% energy loss [1��]. Since formalde-
hyde possesses the same degree of reduction as traditional
sugars, for example, glucose, product yields achieved
from aerobic methane conversion are expected to be
comparable to those of aerobic sugar metabolism. Fur-
thermore, yields of reduced fuels and chemicals will be
limited under aerobic conditions as oxidative phosphor-
ylation competes for reducing equivalents in the form of
NAD(P)H. Scale-up of aerobic methane conversion also
presents a challenge, as methane and oxygen gas transfer
limitations result in poor kinetics. Although these chal-
lenges can be addressed by enhancing the volumetric
mass transfer coefficient (kLa), either from increased gas
flow rate, agitation or improved reactor design, these
improvements result in larger operating and capital
expenses [1��].
Anaerobic methane utilization to producefuels and chemicalsAnaerobic oxidation of methane (AOM) is a significant
biogeochemical process in marine and freshwater sedi-
ments and is important in methane release to the atmo-
sphere (Figure 1) [29,30]. Anaerobic methane oxidizing
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Bioconversion of methane and methanol Bennett et al. 83
Figure 1
syntrophicpartner
MP
methyl-S-CoM
methyl-H4MPT
H4MPT
methylene-H4MPT
methenyl-H4MPT
formyl-H4MPT
formyl-MFR
HS-CoM
F420H2
F420H2
F420
F420H2
F420H2
F420
Fox
Fox
4 Fdred + 2 CO2+ 2 H+
4 Fdox + 2 HS-CoA
Fox
Fox
Fred
Fred
2 Fdred
Fred
Fred
CO2
F420
CH4
F420
CoM-S-S-CoB
CoM-S-S-CoB
CoM-S-S-CoB
2 CoM-S-S-CoB
HS-CoB
2 HS-CoB+
2 HS-CoM
HS-CoM
HS-CoM
HS-CoB
HS-CoA
HS-CoA
acetyl-CoA
acetyl-PO42-
acetate
lactatepyruvate
acetate
Por
Cdh
Mch
Mer
Mtr
HdrDE
Fpo
Rnf
Mtd
Pta
Ack
Acd
HdrABC
HdrABC FrhB
Ftr
Fmd
MvhD
Mcr Hbd
Mhc
Mhc
Nar
MP
H2
MPH2MP
MP
MP
MP
MP
H2
MPH2
MPH2
MQH2
MQ
NO22-
NO32-
ADP+ Pi
2 NADH+ 2H+ 2 NAD+
ADP+ Pi
Pi
ATP
ATP
˜2 Na+
2 Na+
Fe2+
Fe3+
2 H+
2 H+
Current Opinion in Biotechnology
Proposed anaerobic methane oxidation pathways of archaeal methanotrophs and engineered methanotrophic Methanosarcina acetivorans C2A.
Enzymes are shown in bold within rectangles. Enzymes heterologously expressed in M. acetivorans C2A are shown in blue, and pathways utilized
by the engineered M. acetivorans strain to produce acetate and lactate are shown with blue arrows. Enzymes and pathways present only in
ANaerobic MEthanotrophs (ANME) are shown in green. Enzymes and cofactors are as follows: Acd, ADP-forming acetyl-CoA synthetase; Ack,
acetate kinase; Cdh, CO dehydrogenase; Fmd, formylmethylfuran dehydrogenase; Fpo, F420 dehydrogenase (note: this enzyme is replaced by the
homolog Fqo in some ANME); Frh, F420-reducing hydrogenase; Ftr, formylmethanofuran:H4MPT formyltransferase; H4MPT,
tetrahydromethanopterin; Hbd, 3-hydroxylbutyryl-CoA dehydrogenase; HdrABC, soluble heterodisulfide reductase; HdrDE, membrane-bound
heterodisulfide reductase; Mcr, methyl-coenzyme M reducase; Mch, methenyl-H4MPT cyclohydrolase; Mer, methenyl-H4MPT reductase; MF,
methanofuran; Mhc, multiheme cytochrome C; MP, methanophenazine; MQ, methanoquinone (note: some ANME possess MQ instead of MP as
lipophilic electron carrier); Mtd, methenyl-H4MPT; Mtr, methyl-H4MPT:coenzyme M methyltransferase; Mvh, F420-non-reducing hydrogenase; Pta,
phosphotransacetylase; Por, pyruvate ferredoxin oxidoreductase; Rnf, methanophenazine reductase.
www.sciencedirect.com Current Opinion in Biotechnology 2018, 50:81–93
84 Energy biotechnology
archaea, or ANaerobic MEthanotrophs (ANME), were
first discovered obligately associated with bacterial part-
ners that used the reducing equivalents generated during
methane oxidation by ANME to reduce sulfate [30].
ANME belong to four phylogenetic clusters within Eur-
yarchaeota: ANME-1, ANME-2, ANME-3 and GOM Arc
I (formerly ANME-2d) [30]. ANME-1 are related to
Methanosarcinales and Methanomicrobiales whereas others
lie within Methanosarcinales [30]. ANME possess homo-
logs of all methanogenesis enzymes (except for the N5,
N10-methylene-tetrahydromethanopterin reductase
(Mer) in ANME-1), and metatranscriptomic studies of
consortia performing AOM demonstrated transcription of
these enzymes, suggesting methane oxidation to CO2
occurs through a reversal of methanogenesis [31,32].
The key enzyme involved in AOM is a homolog of the
methyl-coenzyme M reductase (MCR), which catalyzes
the formation of methane in methanogenic bacteria (Fig-
ure 1) [33��]. MCRs from ANME and methanogens share
similarity although the MCR from ANME-1 has addi-
tional features, including a tetrapyrrole derivative of the
nickel-containing F430 cofactor and cysteine-rich side
chains [33��].
Despite similarities in MCR structure and sharing
enzymes for AOM, sustained and efficient methane oxi-
dation in methanogens has been challenging. Trace
methane oxidation has been observed in several metha-
nogens [34], and the MCR purified from Methanothermo-bacter marburgensis was found to convert methane and the
heterodisulfide CoM-S-S-CoB into methyl-coenzyme M
(CH3-S-CoM) and coenzyme B (CoB) with rates consis-
tent to in vivo values (Figure 1) [35��]. Additionally, invitro methanol production from CH3-S-CoM was demon-
strated using the Methanosarcina barkeri methanol:coen-
zyme M methyltransferase (MtaABC) [36]. MCRs from
methanogens that catalyze trace methane oxidation share
common features, including four of the five key post-
translational modifications in the active site [37].
Reversal of methanogenesis to oxidize methane requires
a suitable electron acceptor [30]. Removal of reducing
equivalents by syntrophic partner organisms, for example,
sulfate-reducing bacteria, is essential for AOM to be an
energy-yielding process [30]. Studies have demonstrated
direct electron transfer between ANME and bacterial
partners [38,39��]. AOM can also be coupled to the
reduction of nitrate [31], insoluble oxides of Fe3+ and
Mn4+ [40] or soluble electron acceptors such as humic
acids, chelated ferric iron, 9,10-anthraquinone and 2,6-
disulfonate [41��]. The archaeal partner in a consortium
with M. oxyfera, which demonstrated methane oxidation
coupled to denitrification was sequenced, and a draft
genome was assembled with the uncultivated organism
Methanoperedens nitroreducens [7], which contains narGHfor nitrate reduction. Later research reported that Metha-noperedens-like organisms can couple methane oxidation
Current Opinion in Biotechnology 2018, 50:81–93
to particulate Fe3+ and Mn4+ oxides [40], likely mediated
by multiheme cytochrome c enzymes [39��]. Direct elec-
tron transfer between Methanosarcina barkeri [42] or Metha-nosaeta harundinaceae [43] and Geobacter metallireducensduring methanogenesis on ethanol was also demon-
strated. The direct electrical connections observed for
ANME and methanogens suggest that bioreactors for
methane oxidation to fuels and chemicals could utilize
electrochemistry instead of syntrophic partners to remove
reducing equivalents produced during methane
activation.
Currently, no ANME isolate exists, however, engineered
strains can now be envisioned as a number of genetic tools
have been developed for tractable strains including
Methanococcus maripaludis [44] and Methanosarcina species
[45]. These include selection/counter-selection markers,
transformation strategies and replicating vectors for
episomal expression. Recently, Cas9-mediated genome
editing of M. acetivorans was demonstrated utilizing
native homology-dependent repair machinery [46]. The
first report of engineered methane oxidation in a metha-
nogen involved overexpression of ANME-1 MCR in
Methanosarcina acetivorans C2A [47��]. High cell densities
(1010 cells mL�1) of the MCR-expressing strain con-
sumed 15% of supplied methane after 5 days and pro-
duced 10 mM acetate coupled to the reduction Fe3+. Pro-
duction of lactate using the same strain was also reported
[48��], and the yield of 0.59 g g�1 methane was 10-fold
greater than that reported for aerobic production [25��].Additional work engineered an air-adapted strain of M.acetivorans [49] to express the ANME-1 MCR, and it was
cultivated in the anode compartment of a microbial fuel
cell along with Geobacter sulfurreducens and methane-accli-
mated anaerobic digester sludge to produce electricity
[50]. Although methane-fueled microbial fuel cells have
been previously proposed [51], the power density pre-
sented in this work was over twice that obtained previ-
ously [50]. It was postulated that the engineered
M. acetivorans strain converts methane to oxidized inter-
mediates, for example, acetate, which are consumed by
G. sulfurreducens coupled to anode reduction to produce
electricity. Interestingly, all three components of the
consortium were found to be essential for methane
oxidation coupled to electricity production [50], suggest-
ing that anode reduction occurred through electron
shuttles instead of direct contact of a biofilm with the
electrode.
Limitations of AOM coupled to fuel and chemical pro-
duction include slow growth and low energy yields of
anaerobic methanotrophs, resulting in slow AOM rates
[30]. The MCR-expressing M. acetivorans strain also
demonstrated slow growth [47��] despite methane oxida-
tion coupled to Fe3+ reduction being 3.5-times more
energetic (�454 kJ mol�1 [40]) than hydrogenotrophic
methanogenesis (�131 kJ mol�1 [44]) under standard
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Bioconversion of methane and methanol Bennett et al. 85
conditions. All known ANME are related to cytochrome-
containing Methanosarcinales, which possess multiple
routes for energy capture through H+ and Na+ transloca-
tion, and all known ANME genomes encode multiheme
cytochrome c enzymes [30–32]. However, the mode of
energy capture during methane oxidation is unknown,
limiting the ability to elucidate why observed growth
rates for methanogens are orders of magnitude greater
than for ANME or engineered methanogens [30].
As with the aforementioned aerobic scenario, although
anaerobic methane conversion to fuels and chemicals has
been demonstrated, only low yields were achieved at
small scale. However, anaerobic methane conversion does
not suffer from the same limitations that aerobic meth-
ane conversion does. During anaerobic methane conver-
sion, reducing equivalents in the form of NAD(P)H may
be conserved for production of reduced fuels and che-
micals, thus providing the opportunity for improved
product yields from methane. Additionally, gas transfer
limitations are not as critical under anaerobic conditions,
resulting in lower operating and capital expenses as
compared to the aerobic methane conversion scenario.
As a result, anaerobic methane conversion is more ideal
for large scale production of fuels and chemicals. How-
ever, anaerobic methane metabolism is slow compared
with other systems, resulting in low growth rates and
productivities. Therefore, future efforts should be
devoted to improving the rate of anaerobic methane
metabolism for enhanced growth and productivity to
realize scale-up of anaerobic methane conversion to
fuels and chemicals.
Conversion of methane to methanol for use asa substrate for synthetic methylotrophsAlong with the aforementioned biological oxidation of
methane to methanol, chemical conversion of methane to
methanol is also possible. As compared to the biological
oxidation of methane, chemical conversion is faster albeit
suffers from low selectivity and high process demands, for
example, elevated temperatures and pressures [2��]. As a
result, the biological oxidation of methane is more ideal
from an energetics perspective. However, as described
above, several challenges remain before the biological
oxidation of methane can be realized at large scale. In
any case, methanol can serve as an alternative C1 sub-
strate for production of fuels and chemicals. Native
methylotrophs are poor industrial hosts since many are
obligate aerobes and have limited genetic tools, which
are not as well-developed and extensive as those of
platform organisms [2��]. Therefore, there is consider-
able interest in developing synthetic methylotrophs for
the conversion of methanol to fuels and chemicals. In
following sections, we discuss recent advancements
toward achieving synthetic methylotrophy in several
platform organisms.
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Enzyme and pathway considerations forsynthetic methanol utilizationMethanol is first oxidized to formaldehyde via a methanol
oxidoreductase (Figure 2), which include NAD-depen-
dent and PQQ-dependent MDHs from bacteria and
alcohol oxidases (AOXs) from yeast [2��]. NAD-depen-
dent MDHs are ideal for synthetic methylotrophy since
they function aerobically and anaerobically, are expressed
from a single gene and conserve electrons in the form of
NADH [2��,52�].
Formaldehyde is next assimilated via the RuMP pathway,
ribulose bisphosphate (RuBP) pathway or serine cycle
[2��]. Formaldehyde is a toxic intermediate that must be
consumed quickly, and an efficient assimilation pathway
in essential to overcome endogenous formaldehyde dis-
similation [53]. Strategies to eliminate dissimilation
involve gene deletions, for example, formaldehyde dehy-
drogenase ( frmA) in Escherichia coli (Figure 2) [54��].The RuMP pathway is more bioenergetically favorable
than either the serine cycle or RuBP pathway [2��,52�].The two enzymes of the RuMP pathway, hexulose
phosphate synthase (HPS) and phosphohexulose isom-
erase (PHI), fix formaldehyde with the pentose phos-
phate pathway (PPP) intermediate ribulose 5-phosphate
(Ru5P) to generate hexulose 6-phosphate (H6P), which
is isomerized to fructose 6-phosphate (F6P) (Figure 2)
[2��]. Ru5P is a critical intermediate within the RuMP
pathway, and its sustained regeneration is necessary to
sustain methanol assimilation. In principle, only three
heterologous enzymes (MDH, HPS and PHI) are
required to achieve synthetic methylotrophy. However,
recent studies have shown this is not the case and
instead, have shed light on additional limitations,
including poor MDH kinetics and insufficient Ru5P
regeneration.
Sourcing and engineering methanoldehydrogenases (MDHs) for improved kineticpropertiesNAD-dependent MDHs generally exhibit higher affinity
toward higher alcohols, for example, 1-butanol, and meth-
anol oxidation is unfavorable under standard conditions,
explaining why many native methylotrophs are thermo-
philic [2��]. A limited number of NAD-dependent MDHs
have been characterized, notably those from B. methano-licus strains MGA3 and PB1, which each contain three
distinctive MDHs with different kinetic properties [55].
In a recent study, we sourced an NAD-dependent MDH
from the Gram-positive bacterium Bacillus stearothermo-philus [56], which exhibits better reported kinetics than
those from B. methanolicus. This MDH was used to
achieve growth of engineered E. coli on methanol using
a small amount of yeast extract [54��]. The importance of
improved kinetics was demonstrated by realizing that
Mdh2 from B. methanolicus could not achieve methylo-
trophic growth under the same conditions. In both
Current Opinion in Biotechnology 2018, 50:81–93
86 Energy biotechnology
Figure 2
Glucose Methanol
Formaldehyde
G6P GL6P 6PG
F6P
FBP
DHAPDHAP
GAP
GAPE4P
X5P
H6P
R5PRu5P
SBP S7P
hps
tkttkt
fba
fba
pfk
glpX
rpe
mdhfrmA
pgi
fbp
rpi
phi
3PG Ser Gly
Thr
Pyr AcCoA
C1 Pool
CO2
CO2
CO2
KDPG
Biomass, Biofuels & Biochemicals
(-)
Irp
(-)
(+)(+) (-)
Current Opinion in Biotechnology
Strategies to improve synthetic methanol utilization. Methanol assimilation occurs via methanol dehydrogenase (mdh), hexulose phosphate
synthase (hps) and phosphohexulose isomerase ( phi). Formaldehyde dissimilation is eliminated via deletion of formaldehyde dehydrogenase
( frmA). Glucose carbon flux is rerouted through the oxidative pentose phosphate pathway (PPP) for ribulose-5-phosphate (Ru5P) generation via
deletion of phosphoglucose isomerase ( pgi) (shown in green). Heterologous non-oxidative PPP enzymes from Bacillus methanolicus
(phosphofructokinase ( pfk), fructose-bisphosphate aldolase ( fba), transketolase (tkt), ribulose phosphate epimerase (rpe) and sedoheptulose
bisphosphate (glpX)) regenerate Ru5P from fructose-6-phosphate (F6P) (shown in blue). Regulation of endogenous one-carbon (C1) metabolism
via leucine-responsive regulatory protein (lrp) as an alternative route for methanol assimilation (shown in red). Remaining enzymes: fructose
bisphosphatase ( fbp), ribose phosphate isomerase (rpi). Remaining metabolites: glucose-6-phosphate (G6P), 6-phosphogluconolactone (GL6P),
6-phosphogluconate (6PG), ribose-5-phosphate (R5P), fructose bisphosphate (FBP), dihydroxyacetone phosphate (DHAP), glyceraldehyde 3-
phosphate (GAP), xylulose-5-phosphate (X5P), erythrose 4-phosphate (E4P), sedoheptulose bisphosphate (SBP), sedoheptulose-7-phosphate
(S7P), 3-phosphoglycerate (3PG), pyruvate (Pyr), ketodeoxyphosphogluconate (KDPG), serine (Ser), glycine (Gly), threonine (Thr), acetyl-CoA
(AcCoA).
instances, E. coli was engineered to assimilate formalde-
hyde via the RuMP pathway from B. methanolicus. Wu
et al. characterized an NAD-dependent MDH from a
Gram-negative, mesophilic, non-methylotrophic bacte-
rium, Cupriavidus necator (Table 1) [57��]. Directed
molecular evolution was performed to engineer an
MDH variant having improved kinetic properties. Three
mutations were identified (A26V, A31V and A169V), that
when combined, improved methanol affinity and catalytic
efficiency.
Current Opinion in Biotechnology 2018, 50:81–93
Cell-free metabolic engineering todemonstrate and improve methanol utilizationCell-free metabolic engineering has been used to dem-
onstrate methanol conversion and improve MDH kinetic
limitations. Bogorad et al. developed a methanol conden-
sation cycle (MCC) by combining the RuMP pathway
with nonoxidative glycolysis (NOG) for carbon-con-
served, redox-balanced and ATP-independent higher
alcohol production (Table 1) [58�]. Importantly, sugar
phosphates were required to prime MCC, suggesting
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Bioconversion of methane and methanol Bennett et al. 87
Table 1
Strategies and achievements made toward synthetic methanol utilization in recent literature.
Organism Strategy Achievements
Escherichia coli (in vitro)
Scale: mL [58�]� Developed a methanol condensation cycle (MCC) by
combining RuMP and NOG pathways for carbon-
conserved, redox-balanced and ATP-independent
higher alcohol production from methanol and sugar
phosphates in a cell-free system
� MCC produced 13.3 mM ethanol from 33.5 mM
methanol at 80% carbon yield
� MCC produced 2.3 mM 1-butanol from 21.1 mM
methanol at 50% carbon yield
Escherichia coli (in vivo)
Scale: mL [52�]� Performed in silico modeling to demonstrate that
NAD-dependent MDH and RuMP pathway are best
suited for biomass formation from methanol in E. coli
� Characterized multiple recombinant NAD-dependent
MDH and RuMP pathway enzymes in E. coli to identify
best candidates
� Observed up to 39.4% 13C-labeling in glycolytic and
PPP intermediates, specifically hexose 6-phosphates
� Observed RuMP pathway cycling as indicated by
higher-order mass isotopomers
� No growth on methanol was reported
Escherichia coli (in vitro)
[57��]� Identified an activator-independent NAD-dependent
MDH from C. necator N-1, a Gram-negative,
mesophilic, non-methylotrophic bacterium
� Performed directed evolution to improve enzyme
kinetics toward methanol
� Increased methanol affinity (Km) from 132 mM in wild-
type version to 21.6 mM in mutant variant
� Increased methanol catalytic efficiency (Kcat/Km)
from 1.6 M�1 s�1 in wild-type version to 9.3 M�1 s�1 in
mutant variant
� Decreased 1-butanol affinity (Km) from 7.2 mM in
wild-type version to 120 mM in mutant variant
� Decreased 1-butanol catalytic efficiency (Kcat/Km)
from 903 M�1 s�1 in wild-type version to 48 M�1 s�1 in
mutant variant
Escherichia coli (in vitro,
in vivo)
Scale: mL [59��]
� Constructed a scaffoldless enzyme complex of B.
methanolicus Mdh3 and M. gastri Hps-Phi fusion using
an SH3-ligand interaction pair to improve
formaldehyde channeling
� Incorporated E. coli lactate dehydrogenase as an
‘NADH sink’ to improve methanol oxidation kinetics
and carbon flux to F6P
� Improved in vitro F6P production from methanol by
97-fold
� Increased in vivo methanol oxidation rate by 9-fold
and total in vivo methanol consumption by 2.3-fold
Escherichia coli (in vivo)
Scale: mL, L [54��]� Identified a superior NAD-dependent MDH from B.
stearothermophilus to use with the B. methanolicus
RuMP pathway in a DfrmA genetic background
� Supplied small amounts of yeast extract (1 g L�1) as a
co-substrate to stimulate growth on methanol
� Incorporated heterologous pathway for naringenin
production from methanol
� Methanol supplementation improved biomass titers
by up to 50% during growth with a small amount of
yeast extract
� Observed up to 53% 13C-labeling in glycolytic, PPP,
TCA cycle and biomass components, specifically 3PG
� Observed RuMP pathway cycling as indicated by
higher-order mass isotopomers
� Improved naringenin production by 650% over the
empty vector control in methanol and yeast extract
� Observed up to 4.7% 13C-labeling in naringenin and
18% of the total naringenin pool contained at least one
carbon label
Escherichia coli (in vivo)
Scale: mL [69��]� Characterized the native formaldehyde-responsive
promoter (Pfrm) of the formaldehyde dissimilation
operon ( frmRAB) in E. coli
� Performed directed evolution and fluorescence-
activated cell sorting (FACS) in combination with high-
throughput sequencing (Sort-seq) to generate a Pfrm
library
� Developed Pfrm variants that improved formaldehyde
induced expression up to 13-fold and formaldehyde
response up to 3.6-fold
� Achieved autonomous and dynamic regulation of
methylotrophic growth in E. coli via controlling MDH
and RuMP pathway gene expression with native and
engineered Pfrm variants
Escherichia coli (in vivo)
Scale: mL [68]
� Developed a methanol-sensing E. coli strain by
incorporating the MxcQ/MxcE two-component system
from M. organophilum XX
� Constructed a chimeric two-component system by
combining the sensing domain (MxcQ) of M.
organophilum with the transmitter domain (EnvZ) of E.
coli to control the response regulator (OmpR) of E. coli
for activation of the ompC promoter
� Developed a rapid in vivo methanol detection system
in E. coli based on a two-component system from a
native methylotroph
� Achieved a dynamic response range of gene
expression using a range of methanol concentrations
(from 0.01 to 8%)
� Maximum gene expression of ca. 2.5-fold was
achieved with 0.05% methanol
Corynebacterium
glutamicum (in vivo)
Scale: mL, mL [60�]
� Incorporated B. methanolicus MDH and B. subtilis
RuMP pathway into C. glutamicum
� Supplied methanol as a co-substrate in a glucose
minimal medium
� Achieved a methanol consumption rate of
1.7 mM h�1 in a glucose minimal medium
� Methanol supplementation improved biomass titers
by up to 30% during growth in glucose minimal
medium
� Observed up to 25.7% 13C-labeling in M+1 mass
isotopomers of intracellular metabolites, specifically
S7P
www.sciencedirect.com Current Opinion in Biotechnology 2018, 50:81–93
88 Energy biotechnology
Table 1 (Continued )
Organism Strategy Achievements
Corynebacterium
glutamicum (in vivo)
Scale: mL, mL [62��]
� Incorporated B. methanolicus MDH and B. subtilis
RuMP pathway into C. glutamicum for methanol
assimilation
� Supplied methanol as a co-substrate in a glucose or
ribose minimal medium
� Incorporated methanol assimilation pathway into a C.
glutamicum strain capable of non-native cadaverine
production
� Observed up to 25% 13C-labeling in glycolytic and
PPP intermediates, specifically F6P
� Observed RuMP pathway cycling as indicated by
higher-order mass isotopomers
� Observed up to 15.7% 13C-labeling in the non-native,
secreted product cadaverine in ribose minimal
medium
Corynebacterium
glutamicum (in vivo)
Scale: mL [63]
� Performed directed evolution to improve methanol
tolerance of C. glutamicum during growth on glucose
minimal medium
� Performed genome sequencing to identify mutations
responsible for improved methanol tolerance
� Achieved improved growth rates on glucose minimal
medium in the presence of up to 2M methanol
� Identified two point mutations responsible for
improving methanol tolerance (A165T mutation of O-
acetylhomoserine sulfhydrolase MetY and Q342*
mutation leading to a shortened CoA transferase Cat)
Saccharomyces
cerevisiae (in vivo)
Scale: mL [64�]
� Incorporated the methanol assimilation pathway
(AOX, CTA, DAS and DAK) from P. pastoris into S.
cerevisiae
� Supplied small amounts of yeast extract (1 g L�1) as a
co-substrate to stimulate growth on methanol
� Achieved 1.04 g L�1 methanol consumption,
0.26 g L�1 pyruvate production and a 3.13% increase
in biomass titer in methanol minimal medium
� Improved methanol consumption to 2.35 g L�1 and
biomass titer by 11.7% during growth with a small
amount of yeast extract
the importance of sustained Ru5P levels for methanol
utilization. Although methanol conversion was achieved,
productivity decreased after five hours, suggesting insta-
bility of intermediates. It was hypothesized that produc-
tivity and product titers could be improved by achieving
higher fluxes via protein engineering and/or media opti-
mization, and MDH was identified as a critical enzyme for
improvement.
Price et al. constructed a scaffoldless enzyme complex
composed of Mdh3 from B. methanolicus and an HPS-PHI
fusion protein from Mycobacterium gastri (Table 1) [59��].This complex promoted efficient formaldehyde channel-
ing to improve carbon flux from methanol to F6P. An
‘NADH sink’ was also developed by incorporating the
LDH from E. coli, which catalyzes the NADH-dependent
reduction of pyruvate to lactate, to scavenge the NADH
from methanol oxidation, preventing formaldehyde
reduction. The complex with LDH improved in vitroF6P production by 97-fold compared to unassembled
enzymes, and improvements were also realized in vivoas the complex increased methanol uptake rate by 9-fold
and total methanol consumption by 2.3-fold compared to
unassembled enzymes. The discrepancy between in vitroand in vivo improvements highlights the difficulties with
engineering complex biological systems. When transition-
ing from cell-free to in vivo conditions, additional biological
factors must be considered, for example, gene regulation,
that may be assumed negligible during in vitro studies.
Engineering Escherichia coli to assimilatemethanol for in vivo growth and metaboliteproductionMuller et al. reported in vivo 13C-methanol assimilation
in E. coli via incorporation of Mdh2, HPS and PHI from
Current Opinion in Biotechnology 2018, 50:81–93
B. methanolicus (Table 1) [52�]. The engineered E. coliexhibited up to 39.4% 13C-labeling in glycolytic
and PPP intermediates. RuMP pathway cycling
was demonstrated as higher-order mass isotopomers
were observed. Although methanol assimilation was
achieved, no growth on methanol was reported,
suggesting limitations downstream of methanol
oxidation.
Whitaker et al. reported methylotrophic growth of
engineered E. coli by sourcing a superior MDH from
B. stearothermophilus, which was expressed with the
RuMP pathway from B. methanolicus in a DfrmA back-
ground (Table 1) [54��]. Methylotrophic growth was
achieved with a small amount of yeast extract. Upon
yeast-extract exhaustion, growth was sustained on
methanol for ca. 64 h, during which time ca. 10 mM
methanol was consumed at a rate of 19 mg gDW�1 h�1,
significantly less than that of native methylotrophs,
further suggesting limitations downstream of methanol
oxidation. Methanol supplementation improved bio-
mass titers by nearly 50% in bioreactors, during which
time a biomass yield of 0.344 gDW g�1 methanol was
achieved, comparable to that of native methylotrophs.
Up to 53% 13C-labeling was observed in glycolytic, PPP
and tricarboxylic acid cycle intermediates, as well as
hydrolyzed biomass components. RuMP pathway
cycling was also demonstrated as higher-order mass
isotopomers were observed. By incorporating the nar-
ingenin pathway into engineered E. coli, naringenin
production was improved 650% over the empty vector
control in 13C-methanol and yeast extract. Up to 4.7%
average 13C-labeling was observed in naringenin with
18% of the total naringenin pool containing at least one
carbon label.
www.sciencedirect.com
Bioconversion of methane and methanol Bennett et al. 89
Other synthetic methylotrophs for in vivomethanol assimilation and metaboliteproductionWitthoff et al. demonstrated in vivo methanol assimila-
tion in C. glutamicum using a similar strategy as those
used in E. coli (Table 1) [60�]. The engineered strain
exhibited a methanol uptake rate of 1.7 mM h�1 and a
30% improvement in biomass titer in glucose minimal
medium. Up to 25.7% 13C-labeling in M+1 mass iso-
topomers was observed in intracellular metabolites using
a formaldehyde dissimilation deficient strain, con-
structed via deletion of acetaldehyde dehydrogenase
(ald) and mycothiol-dependent formaldehyde dehydro-
genase (adhE) [61]. Lebmeier et al. engineered C. glu-tamicum to convert 13C-methanol into the non-native,
secreted product cadaverine using a similar strategy
(Table 1) [62��]. Up to 15.7% 13C-labeling in cadaverine
was observed in ribose minimal medium. C. glutamicumwas also evolved for improved methanol tolerance
(Table 1) [63].
Dai et al. demonstrated in vivo methanol assimilation in
the non-methylotrophic yeast Saccharomyces cerevisiae by
integrating AOX, catalase (CAT), dihydroxyacetone
synthase (DAS) and dihydroxyacetone kinase (DAK),
all from Pichia pastoris, into the chromosome (Table 1)
[64�]. DAS and DAK compose the xylulose monopho-
sphate (XuMP) pathway for formaldehyde assimilation
[65]. The engineered strain consumed 1.04 g L�1 meth-
anol, produced 0.26 g L�1 pyruvate and exhibited a
3.13% improvement in biomass titer in methanol mini-
mal medium. Yeast extract supplementation improved
methanol consumption to 2.35 g L�1 and biomass titer
by 11.7%, consistent with previous findings in E. coli[54��].
Developing methanol and formaldehyderesponsiveness in synthetic methylotrophsOne limitation of synthetic methylotrophy is the inabil-
ity to regulate gene expression in response to methanol
and/or formaldehyde, which leads to reduced gene
expression and metabolic activity during growth on
methanol [66]. Native methylotrophs regulate gene
expression via methanol-responsive and/or formalde-
hyde-responsive promoters or systems [67,68]. Upregu-
lation of RuMP pathway and PPP genes in B. methano-licus during methylotrophic growth improves methanol
tolerance and uptake rate [67]. Regulating gene expres-
sion in response to methanol is a critical component for
synthetic methylotrophy.
Rohlhill et al. demonstrated methanol-responsiveness in
E. coli by refactoring mdh, hps and phi expression with the
native formaldehyde-responsive promoter (Pfrm) of the
formaldehyde dissimilation operon ( frmRAB) in E. coli(Figure 3, Table 1) [69��]. Directed evolution and Sort-
Seq were performed to construct a Pfrm library, resulting
www.sciencedirect.com
in variants with improved promoter activity. Methylo-
trophic growth of engineered E. coli was achieved with a
small amount of yeast extract when mdh, hps and phi were
autonomously and dynamically regulated using native
and engineered Pfrm variants.
Selvamani et al. developed a methanol-sensing E. colistrain by combining the sensing domain (MxcQ) of
Methylobacterium organophilum with the transmitter
domain (EnvZ) of E. coli to control the response regulator
(OmpR) of E. coli for activation of the ompC promoter
(Figure 3, Table 1) [68]. This resulted in a rapid in vivomethanol detection system that exhibited a dynamic
range of gene expression in response to a wide range
of methanol concentrations (0.01–8%). ompC gene expres-
sion was improved a maximum of ca. 2.5-fold during
exposure to 0.05% methanol. Ganesh et al. reported a
similar strategy that combined the sensing domain
(MxaY) of Paracoccus denitrificans with the transmitter
domain (EnvZ) of E. coli [70].
Strategies to improve ribulose 5-phosphate(Ru5P) (re)generationA critical limitation of synthetic methylotrophy is ineffi-
cient Ru5P regeneration. One strategy to improve Ru5P
regeneration involves refactoring the expression of native
PPP genes using native or engineered Pfrm promoters or a
methanol-sensing system (Figure 3) [68,69��], which
would upregulate gene expression during growth on
methanol, emulating native methylotrophs and providing
sufficient flux for Ru5P regeneration. This strategy is
readily applicable to other target genes as well that
may be later identified as important for synthetic
methylotrophy.
Another strategy to improve Ru5P regeneration is via
overexpression of heterologous PPP enzymes. B. metha-nolicus contains plasmid homologs of chromosomal non-
oxidative PPP enzymes that have evolved to favor the
production of Ru5P from F6P for methanol assimilation
(Figure 2) [71]. Overexpression of heterologous, methy-
lotrophic PPP enzymes improves the kinetics and favor-
ability of Ru5P regeneration for improved methanol
utilization in E. coli [66]. Expression of these genes could
also be refactored for methanol-responsiveness and/or
formaldehyde-responsiveness.
Another strategy to improve Ru5P generation involves
deletion of phosphoglucose isomerase ( pgi). Glycolysis is
the primary route for glucose catabolism in wild-type
E. coli, limiting carbon flux through the PPP [66]. Dele-
tion of pgi reroutes glucose carbon flux through the
oxidative PPP for sustained Ru5P generation and
improves methanol utilization in E. coli (Figure 2) [66].
Furthermore, deletion of pgi acts to conserve methanol
carbon since F6P cannot be metabolized via decarboxyl-
ation reactions in the oxidative PPP.
Current Opinion in Biotechnology 2018, 50:81–93
90 Energy biotechnology
Figure 3
Methanol Formaldehyde
pfk
tktrpe
hpsmdh phi
fba glpX
ChromosomalExpression
Episomal Expression
MDH
EnvZMxc
Q
Om
pRPompC
PompC
Pfrm
PP
Current Opinion in Biotechnology
Strategies to regulate gene expression in response to methanol and formaldehyde. When the methanol-sensing domain (MxcQ) of
Methylobacterium organophilum is fused with the transmitter domain (EnvZ) of E. coli, the response regulator (OmpR) of E. coli activates the
ompC promoter (PompC). Upon methanol oxidation via MDH, formaldehyde activates the formaldehyde-responsive promoter (Pfrm) of E. coli. Genes
listed serve as examples: methanol dehydrogenase (mdh), hexulose phosphate synthase (hps), phosphohexulose isomerase ( phi),
phosphofructokinase ( pfk), fructose-bisphosphate aldolase ( fba), transketolase (tkt), ribulose phosphate epimerase (rpe) and sedoheptulose
bisphosphate (glpX).
Exploring amino acid metabolism to improvesynthetic methanol assimilationSince yeast extract, which is primarily composed of amino
acids, stimulates synthetic methylotrophy, the metabo-
lism of all 20 amino acids and the regulatory networks in
which they are involved were examined [72]. It was
determined that co-utilization of threonine leads to
improved methanol assimilation in a synthetic E. colimethylotroph, which resulted from activation of endoge-
nous C1 metabolism via high flux from threonine to
glycine to serine under threonine growth conditions
(Figure 2) [72]. To verify the phenotype, a global regula-
tor that regulates this pathway was identified and exam-
ined. This regulator, the leucine-responsive regulatory
protein (Lrp), represses threonine dehydrogenase and
serine hydroxymethyltransferase, which respectively cat-
alyze the conversion of threonine to glycine and glycine
to serine. For improved methanol assimilation, these
pathways should be active. Therefore, the lrp gene was
deleted in a synthetic E. coli methylotroph, which
resulted in improved growth and methanol assimilation
compared to the lrp-intact strain [72]. This study provides
the basis for exploring other regulatory networks that
Current Opinion in Biotechnology 2018, 50:81–93
would further enhance synthetic methylotrophy and a
strain capable of growth solely on methanol.
Future perspectives and recommendationsTwo key limitations were identified while attempting to
engineer synthetic methylotrophs for methanol utiliza-
tion. First, methanol oxidation is limited by MDH kinet-
ics. Sourcing alternative or engineering current MDHs
for improved kinetics can overcome this limitation. Sec-
ond, methanol assimilation is limited by inefficient
Ru5P regeneration. Several strategies can overcome this,
including refactoring native PPP gene expression for
methanol-responsiveness, incorporating a heterologous
non-oxidative PPP with improved F6P to Ru5P kinetics
or deleting pgi to reroute glucose flux entirely toward
Ru5P.
Since growth on methanol as the sole carbon source has
yet been achieved, future studies should explore various
cellular mechanisms, not just those limited to methanol
oxidation and Ru5P regeneration. For example, transcrip-
tomic approaches may identify gene(s) that are indirectly
involved in methanol metabolism but could prove
www.sciencedirect.com
Bioconversion of methane and methanol Bennett et al. 91
essential for achieving growth on methanol. Another
example is to evolve current methylotrophic phenotypes
for improved properties and identify the mutations
responsible, especially if autonomous growth on metha-
nol as the sole carbon source is achieved. Another
approach is to screen genomic or enriched metagenomic
libraries from native methylotrophs under conditions that
assess the impact of multiple genes combinatorially
[73,74]. Endogenous C1 metabolism could also be
explored as an alternative route for methanol utilization
(Figure 2) [72]. Since methanol likely causes carbon
starvation responses in non-methylotrophic organisms,
these mechanisms could be explored as well.
Acknowledgement
Financial support from ARPA-E through contract no. DE-AR0000432 isgratefully acknowledged.
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