engineering the bioconversion of methane and methanol to...

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

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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


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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


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


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


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


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



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.


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



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

(-)

(+)(+) (-)


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.


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



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


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



� 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


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


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


Table 1 (Continued )

Organism Strategy Achievements

Corynebacterium


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 up to 15.7% 13C-labeling in the non-native,

secreted product cadaverine in ribose minimal

medium

Corynebacterium


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


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.



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


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.



Figure 3

Methanol Formaldehyde

pfk

tktrpe

hpsmdh phi

fba glpX

ChromosomalExpression

Episomal Expression

MDH

EnvZMxc

Q

Om

pRPompC

PompC

Pfrm

PP


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


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



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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First reported demonstration of in vivo methanol assimilation in E. colialthough growth on methanol was not reported.

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First reported demonstration of growth on methanol and in vivo methanolbioconversion to naringenin in E. coli.

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Performed directed evolution to evolve an MDH from C. necator forimproved kinetics toward methanol.

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First demonstration of a methanol condensation cycle for in vitro higheralcohol production from methanol.

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First demonstration of a scaffoldless enzyme complex that improvesmethanol utilization in vitro and in vivo.

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First reported demonstration of in vivo methanol assimilation in C.glutamicum.

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First reported demonstration of in vivo methanol bioconversion to cada-verine in C. glutamicum.

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67. Jakobsen OM, Benichou A, Flickinger MC, Valla S, Ellingsen TE,Brautaset T: Upregulated transcription of plasmidand chromosomal ribulose monophosphate pathwaygenes is critical for methanol assimilation rate andmethanol tolerance in the methylotrophicbacterium Bacillus methanolicus. J Bacteriol 2006, 188:3063-3072.

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Rohlhill J, Sandoval NR, Papoutsakis ET: Sort-seq approach toengineering a formaldehyde-inducible promoter fordynamically regulated Escherichia coli growth on methanol.ACS Synth Biol 2017, 6:1584-1595.

First reported demonstration of autonomously-regulated growth onmethanol in E. coli using a native formaldehyde-responsive promoter.

70. Ganesh I, Vidhya S, Eom GT, Hong SH: Construction ofmethanol-sensing Escherichia coli by the introduction of aParacoccus denitrificans MxaY-based chimeric two-component system. J Microbiol Biotechnol 2017, 27:1106-1111.

71. Muller JE, Heggeset TM, Wendisch VF, Vorholt JA, Brautaset T:Methylotrophy in the thermophilic Bacillus methanolicus,basic insights and application for commodity production frommethanol. Appl Microbiol Biotechnol 2015, 99:535-551.

72. Gonzalez JE, Bennett RK, Papoutsakis ET, Antoniewicz MR:Methanol assimilation in Escherichia coli is improved by co-utilization of threonine and deletion of leucine-responsiveregulatory protein. Metab Eng 2017 http://dx.doi.org/10.1016/j.ymben.2017.11.015.

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