Extracellular fungal polyol lipids A new class of...
18
Contents lists available at ScienceDirect Biotechnology Advances journal homepage: www.elsevier.com/locate/biotechadv Research review paper Extracellular fungal polyol lipids: A new class of potential high value lipids Luis A. Garay a , Irnayuli R. Sitepu a,b , Tomas Cajka c , Jian Xu d , Hui Ean Teh e , J. Bruce German d , Zhongli Pan e,f , Stephanie R. Dungan d,g , David E. Block h,i , Kyria L. Boundy-Mills a, ⁎ a Phaff Yeast Culture Collection, Department of Food Science and Technology, University of California, One Shields Ave, Davis, CA 95616, USA b Biotechnology Department, Indonesia International Institute for Life Sciences (i3L), Jalan Pulo Mas Barat Kav. 88, East Jakarta, DKI Jakarta 13210, Indonesia c UC Davis Genome Center—Metabolomics, University of California Davis, 451 Health Sciences Drive, Davis, CA 95616, USA d Department of Food Science and Technology, University of California Davis, One Shields Ave, Davis, CA 95616, USA e Department of Biological and Agricultural Engineering, University of California Davis, One Shields Ave, Davis, CA 95616, USA f Healthy Processed Foods Research Unit, Western Regional Research Center, USDA, Agricultural Research Service, 800 Buchanan Street, Albany, CA 94710, USA g Department of Chemical Engineering, University of California Davis, One Shields Ave, Davis, CA 95616, USA h Department of Viticulture and Enology, University of California Davis, One Shields Avenue, Davis, CA 95616, USA i Chemical Engineering and Material Science, University of California Davis, One Shields Avenue, Davis, CA 95616, USA ARTICLE INFO Keywords: Liamocins Polyol esters of fatty acids Extracellular fungal glycolipid biosurfactants Aureobasidium Rhodotorula Biorefinery ABSTRACT Extracellular fungal glycolipid biosurfactants have attracted attention because productivities can be high, cheap substrates can be used, the molecules are secreted into the medium and the downstream processing is relatively simple. Three classes of extracellular fungal glycolipid biosurfactants have provided most of the scientific ad- vances in this area, namely sophorolipids, mannosylerythritol lipids and cellobioselipids. Polyol lipids, a fourth class of extracellular fungal glycolipid biosurfactants, comprise two groups of molecules: liamocins produced by the yeast-like fungus Aureobasidium pullulans, and polyol esters of fatty acids, produced by some Rhodotorula yeast species. Both are amphiphilic, surface active molecules with potential for commercial development as surfactants for industrial and household applications. The current knowledge of polyol lipids highlights an emerging group of extracellular fungal glycolipid biosurfactants and provides a perspective of what next steps are needed to harness the benefits and applications of this novel group of molecules. 1. Introduction Polyol lipids are a class of extracellular fungal glycolipid bio- surfactants (EFGB) that differ from other classes of EFGB such as so- phorolipids (SL) (Van Bogaert et al., 2011, 2007b), mannosylerythritol lipids (MEL) (Kitamoto et al., 1993) and cellobiose lipids (Puchkov et al., 2002) in that no saccharide moiety is present in their structures: the polar moiety is a polyol. They can be divided into two groups: liamocins and polyol esters of fatty acids (PEFA). These are the only currently known groups of EFGB that have a polyol head group. Polyol lipids are a less studied class of biosurfactants made by yeast and yeast- like fungi that hold promise for commercialization. Polyol lipid-se- creting fungi are also an emerging topic for basic research on secretion systems and secondary metabolite biosynthesis. Unlike extensively studied SL, polyol lipids contain polyol moieties rather than carbohy- drates as the hydrophilic head groups. Polyol lipid biosynthesis and secretion have been reported by several species of the Ascomycete polymorphic yeast-like fungus Aureobasidium, and by six different Basidiomycete yeast species within the Sporidiobolales order, all of them belonging to the polyphyletic Rhodotorula clade (see Table 1). This discussion will focus first on liamocins, then on PEFA, comparing and contrasting these groups, and highlighting where further research is needed. Polyol lipids produced chemically by enzymatic methods are beyond the scope of the present discussion. However, the reader is encouraged to see the work of Ducret et al. (1996) and Lortie (1997) for a deeper understanding of enzymatic synthesis of polyol lipids. 2. Liamocins 2.1. Producing strains Liamocin production was first reported by Ruinen and Deinema (1964) by two strains, numbers 30 and 272, both described as Pullularia pullulans (currently known as Aureobasidium pullulans), collected by Deinema from the phyllosphere of Java in Indonesia and Suriname. These excreted material contained hydroxylated fatty acids and a C 6 - https://doi.org/10.1016/j.biotechadv.2018.01.003 Received 22 May 2017; Received in revised form 7 December 2017; Accepted 3 January 2018 ⁎ Corresponding author at: One Shields Avenue, Davis, CA 95616-8598, USA. E-mail addresses: [email protected] (L.A. Garay), [email protected] (I.R. Sitepu), [email protected] (T. Cajka), [email protected] (J. Xu), [email protected] (H.E. Teh), [email protected] (Z. Pan), [email protected] (S.R. Dungan), [email protected] (D.E. Block), [email protected] (K.L. Boundy-Mills). Biotechnology Advances 36 (2018) 397–414 Available online 05 January 2018 0734-9750/ © 2018 Elsevier Inc. All rights reserved. T
Transcript of Extracellular fungal polyol lipids A new class of...
Contents lists available at ScienceDirect
Biotechnology Advances
journal homepage: www.elsevier.com/locate/biotechadv
Research review paper
Extracellular fungal polyol lipids: A new class of potential high value lipids
Luis A. Garaya, Irnayuli R. Sitepua,b, Tomas Cajkac, Jian Xud, Hui Ean Tehe, J. Bruce Germand,Zhongli Pane,f, Stephanie R. Dungand,g, David E. Blockh,i, Kyria L. Boundy-Millsa,⁎
a Phaff Yeast Culture Collection, Department of Food Science and Technology, University of California, One Shields Ave, Davis, CA 95616, USAb Biotechnology Department, Indonesia International Institute for Life Sciences (i3L), Jalan Pulo Mas Barat Kav. 88, East Jakarta, DKI Jakarta 13210, IndonesiacUC Davis Genome Center—Metabolomics, University of California Davis, 451 Health Sciences Drive, Davis, CA 95616, USAd Department of Food Science and Technology, University of California Davis, One Shields Ave, Davis, CA 95616, USAe Department of Biological and Agricultural Engineering, University of California Davis, One Shields Ave, Davis, CA 95616, USAfHealthy Processed Foods Research Unit, Western Regional Research Center, USDA, Agricultural Research Service, 800 Buchanan Street, Albany, CA 94710, USAg Department of Chemical Engineering, University of California Davis, One Shields Ave, Davis, CA 95616, USAhDepartment of Viticulture and Enology, University of California Davis, One Shields Avenue, Davis, CA 95616, USAi Chemical Engineering and Material Science, University of California Davis, One Shields Avenue, Davis, CA 95616, USA
A R T I C L E I N F O
Keywords:LiamocinsPolyol esters of fatty acidsExtracellular fungal glycolipid biosurfactantsAureobasidiumRhodotorulaBiorefinery
A B S T R A C T
Extracellular fungal glycolipid biosurfactants have attracted attention because productivities can be high, cheapsubstrates can be used, the molecules are secreted into the medium and the downstream processing is relativelysimple. Three classes of extracellular fungal glycolipid biosurfactants have provided most of the scientific ad-vances in this area, namely sophorolipids, mannosylerythritol lipids and cellobioselipids. Polyol lipids, a fourthclass of extracellular fungal glycolipid biosurfactants, comprise two groups of molecules: liamocins produced bythe yeast-like fungus Aureobasidium pullulans, and polyol esters of fatty acids, produced by some Rhodotorulayeast species. Both are amphiphilic, surface active molecules with potential for commercial development assurfactants for industrial and household applications. The current knowledge of polyol lipids highlights anemerging group of extracellular fungal glycolipid biosurfactants and provides a perspective of what next stepsare needed to harness the benefits and applications of this novel group of molecules.
1. Introduction
Polyol lipids are a class of extracellular fungal glycolipid bio-surfactants (EFGB) that differ from other classes of EFGB such as so-phorolipids (SL) (Van Bogaert et al., 2011, 2007b), mannosylerythritollipids (MEL) (Kitamoto et al., 1993) and cellobiose lipids (Puchkovet al., 2002) in that no saccharide moiety is present in their structures:the polar moiety is a polyol. They can be divided into two groups:liamocins and polyol esters of fatty acids (PEFA). These are the onlycurrently known groups of EFGB that have a polyol head group. Polyollipids are a less studied class of biosurfactants made by yeast and yeast-like fungi that hold promise for commercialization. Polyol lipid-se-creting fungi are also an emerging topic for basic research on secretionsystems and secondary metabolite biosynthesis. Unlike extensivelystudied SL, polyol lipids contain polyol moieties rather than carbohy-drates as the hydrophilic head groups. Polyol lipid biosynthesis andsecretion have been reported by several species of the Ascomycetepolymorphic yeast-like fungus Aureobasidium, and by six different
Basidiomycete yeast species within the Sporidiobolales order, all ofthem belonging to the polyphyletic Rhodotorula clade (see Table 1).This discussion will focus first on liamocins, then on PEFA, comparingand contrasting these groups, and highlighting where further researchis needed. Polyol lipids produced chemically by enzymatic methods arebeyond the scope of the present discussion. However, the reader isencouraged to see the work of Ducret et al. (1996) and Lortie (1997) fora deeper understanding of enzymatic synthesis of polyol lipids.
2. Liamocins
2.1. Producing strains
Liamocin production was first reported by Ruinen and Deinema(1964) by two strains, numbers 30 and 272, both described as Pullulariapullulans (currently known as Aureobasidium pullulans), collected byDeinema from the phyllosphere of Java in Indonesia and Suriname.These excreted material contained hydroxylated fatty acids and a C6-
https://doi.org/10.1016/j.biotechadv.2018.01.003Received 22 May 2017; Received in revised form 7 December 2017; Accepted 3 January 2018
⁎ Corresponding author at: One Shields Avenue, Davis, CA 95616-8598, USA.E-mail addresses: [email protected] (L.A. Garay), [email protected] (I.R. Sitepu), [email protected] (T. Cajka), [email protected] (J. Xu), [email protected] (H.E. Teh),
[email protected] (Z. Pan), [email protected] (S.R. Dungan), [email protected] (D.E. Block), [email protected] (K.L. Boundy-Mills).
Biotechnology Advances 36 (2018) 397–414
Available online 05 January 20180734-9750/ © 2018 Elsevier Inc. All rights reserved.
T
Table1
Ascom
ycetou
san
dBa
sidiom
ycetou
sspecieskn
ownto
synthe
size
andsecretepo
lyol
lipidsin
yields
equa
lorab
ove1g/
L.Abb
reviations:A
TCC:A
merican
Type
Culture
Collection,
Virginia,
USA
;CBS
:Cen
traa
lbureauvo
orSchimmelcu
ltures,T
heNethe
rlan
ds;NRRL:
USD
A-ARSCulture
Collection,
Illin
ois,
USA
.
Species
Strain
IDnu
mbe
rin
othe
rco
llections
Yearof
publication
Source
ofthestrain
Referen
ce
Ascom
ycota(liamoc
ins)
Aureoba
sidium
sp.
A-91
1992
Unk
nown
Nag
ataet
al.(19
93);Ta
buch
ian
dKay
ano(199
2)Aureoba
sidium
sp.
A-2,
1994
Amixture
ofox
idativeye
astfrom
naturalsources,
location
unkn
own
Kurosaw
aet
al.(19
94)
Aureoba
sidium
sp.
A-21M
1994
Aureoba
sidium
pullu
lans
272
1964
Phyllosphe
reof
trop
ical
folia
ge,J
ava
Ruine
nan
dDeine
ma(196
4)Aureoba
sidium
pullu
lans
IFO
7757
1992
Unk
nown
Tabu
chian
dKay
ano(199
2)Aureoba
sidium
pullu
lans
CU
2NRRL58
515
2016
Lagerstroemia
loud
onii,
Bang
kok,
Thailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
4NRRL58
517
2016
Cresentia
alata,
Patumthan
i,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
10NRRL58
523
2016
Ficusbenjam
inaL.,U
bonratch
atha
ni,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
12NRRL58
525
2016
Tamarindu
sindica
L.,U
bonratch
atha
ni,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
13NRRL58
526
2016
Tamarindu
sindica
L.,U
bonratch
atha
ni,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
16NRRL58
529
2016
Ficusbenjam
inaL.,U
bonratch
atha
ni,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
22NRRL58
535
2012
Garciniaman
gostan
aL.,C
hantha
buri,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
23NRRL58
536
2016
Eugeniajunb
os,C
hantha
buri,T
haila
ndMan
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
27NRRL58
540
2016
Kerriod
oxaelegan
s,Prachu
apkh
irikha
nTh
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
28NRRL58
541
2016
Pinu
smerku
sii,Prachu
apkh
irikha
n,Th
ailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
31NRRL58
544
2009
,20
11Pinu
smerku
sii(Needlepine
),Prachu
apkh
irikha
n,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.
(201
1)Aureoba
sidium
pullu
lans
CU
35NRRL58
547
2009
,20
11Hopea
ferrea
Lane
ss.,Pa
thum
than
i,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.
(201
1)Aureoba
sidium
pullu
lans
CU
39NRRL58
551
2009
,20
11Mim
usopselengi
L.(Spa
nish
cherry),Non
gkha
i,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.
(201
1)Aureoba
sidium
pullu
lans
CU
40NRRL58
552
2016
Sand
oricum
indicum
Cav
.,Chian
gMai,T
haila
ndMan
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
41NRRL58
553
2016
Syzygium
cuminiL
.,Ray
ong,
Thailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
42NRRL58
554
2016
Man
gifera
indica
L,Ray
ong,
Thailand
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
43NRRL50
380
2011
,20
13Le
afof
CassiafistulaL.
(golde
nshow
ertree),
Udo
ntha
ni,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.
(201
1);P
rice
etal.(20
13)
Aureoba
sidium
pullu
lans
CU
44NRRL58
555
2012
Saraca
indica,R
atch
aburi,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.,
2012
Aureoba
sidium
pullu
lans
CU
45NRRL58
556
2009
,20
11Sa
man
easaman
(EastIndian
walnu
t)Jacq
.,Ba
ngko
k,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.
(201
1)Aureoba
sidium
pullu
lans
CU
46NRRL58
557
2016
Man
gifera
indica
L.,T
rang
,Tha
iland
Man
itch
otpisitet
al.(20
09)
Aureoba
sidium
pullu
lans
CU
47NRRL58
558
2009
,20
11Ta
marindu
sindica
L.(Tam
arind),P
atum
than
i,Th
ailand
Man
itch
otpisitet
al.(20
09);Man
itch
otpisitet
al.
(201
1)Aureoba
sidium
pullu
lans
NRRLY-258
120
11Unk
nown
Man
itch
otpisitet
al.(20
11)
Aureoba
sidium
pullu
lans
NRRLY-402
620
11Unk
own
Man
itch
otpisitet
al.(20
11)
Aureoba
sidium
pullu
lans
NRRLY-458
820
11Pitchwou
ndon
pine
,loc
ationun
know
nMan
itch
otpisitet
al.(20
11)
Aureoba
sidium
pullu
lans
NRRLY-129
7420
11,20
12Se
aweed,
Florida,
USA
Man
itch
otpisitet
al.(20
11);Man
itch
otpisitet
al.
(201
2)Aureoba
sidium
pullu
lans
RSU
6NRRL50
383
2015
Leaf
inRey
kjav
ik,Iceland
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
9NRRL62
031
2016
Unide
ntified
leaf,N
akornratch
asim
a,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
12NRRL50
381
2015
Leaf,C
hiya
phum
,Tha
iland
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
13NRRL62
034
2014
Leaf,C
honb
uri,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
15NRRL62
036
2012
Leaf,R
atch
aburi,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
17NRRL62
038
2014
Leaf,P
atalun
g,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
18NRRL62
039
2014
Leaf,C
honb
uri,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
19NRRL62
040
2016
Unidentified
leaf,P
atalun
g,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
20NRRL62
041
2014
Leaf,P
atalun
g,Th
ailand
Man
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
21NRRL62
042
2016
Unidentified
leaf,P
atalun
gMan
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
29NRRL50
384
2015
Man
gifera
indica
L.,B
angk
ok,T
haila
ndMan
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
RSU
32NRRL50
382
2015
Leaf,B
angk
ok,T
haila
ndMan
itch
otpisitet
al.(20
12)
Aureoba
sidium
pullu
lans
L3-G
PY20
15Liliu
mlancifo
lium
Thun
b(Lily
Wild
Flow
er),
Kim
etal.(20
15)
(con
tinuedon
next
page)
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
398
Table1(con
tinued)
Species
Strain
IDnu
mbe
rin
othe
rco
llections
Yearof
publication
Source
ofthestrain
Referen
ce
Aureoba
sidium
pullu
lans
P520
15Man
grov
esystem
sin
China
Liet
al.(20
15)
Aureoba
sidium
man
sonii
IFO
9233
1992
Unk
nown
Tabu
chian
dKay
ano(199
2)Aureoba
sidium
microstictum
IFO
3206
919
92Unk
nown
Tabu
chian
dKay
ano(199
2)
Basidiom
ycota(PEF
A)
Rho
dosporidiobolusazoricus
(previou
slycalle
dRho
dotorula
glutinis)
CBS
4648
1964
Leaf
surfaceof
Cacao
plan
ts,Gha
naRuine
nan
dDeine
ma(196
4);T
ulloch
andSp
encer
(196
4)Rho
dotorula
glutinis
CBS
3044
1964
Leaf
ofDesmod
ium
repens,inho
tho
use,
Wag
eninge
n,Nethe
rlan
dsRuine
nan
dDeine
ma(196
4)
Rho
dotorula
glutinis
16A8
1964
Flow
ers,
Can
ada
Tullo
chan
dSp
encer(196
4)Rho
dotorula
toruloides
(previou
slycalle
dRho
dotorula
glutinis)
IIP-30
ATC
C20
4091
1992
Hyd
rocarbon
contam
inated
soil,
India
John
sonet
al.(19
92)
Rho
dotorula
gram
inis
6CB
1961
Flow
ers,
Can
ada
Tullo
chan
dSp
encer(196
4)Rho
dotorula
gram
inis
UCDFS
T05
-503
2016
Oliv
efly,
California,
USA
Garay
etal.(20
17b)
Rho
dotorula
gram
inis
CBS
3043
1961
,19
64Le
afsurfaceof
citrus
plan
ts,Bo
gor,
Indo
nesia
Deine
ma(196
1);R
uine
n(196
3)Rho
dotorula
gram
inis
CBS
2826
2K53
1958
,19
61Grass,N
orth
Island
,New
Zealan
dDeine
ma(196
1);d
iMen
na(195
8)Rho
dotorula
babjevae
UCDFS
T04
-877
2016
Oliv
efly,
California,
USA
Cajka
etal.(20
16)
Rho
dotorula
babjevae
UCDFS
T04
-830
2016
Maleolivefly,
California,
USA
Garay
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T05
-613
2016
Oliv
etree,C
alifornia,
USA
Garay
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T05
-736
2016
Femaleolivefly,
California,
USA
Garay
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T05
-775
2016
Dry
sapof
olivetree,C
alifornia,
USA
Garay
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T06
-542
2016
Oliv
e-flyinfested
olive,
California,
USA
Garay
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T67
-102
2016
Seawater,C
alifornia,
USA
Garay
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T67
-436
2016
Exud
ateof
Pterocarya
rhoifolia
,Kaida
,Kiso,
Japa
nGaray
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T67
-458
2016
Exud
ateof
Betula
erman
i,Mt.Fu
ji,Japa
nGaray
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T67
-478
2016
Exud
ateof
Prun
ussargentii,J
atan
i,Yam
agata,
Japa
nGaray
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T67
-506
2016
Exud
ateof
Ulm
usda
vidian
ava
r.japonica,H
ibara
(Fuk
ushima),Japa
nGaray
etal.(20
17b)
Rho
dotorula
babjevae
UCDFS
T68
-916
.120
16Insect
frassin
Alnus
sp.(alde
r)tree,B
ritish
Colum
bia,
Can
ada
Garay
etal.(20
17b)
Rho
dotorula
diobovata
UCDFS
T08
-225
2016
Seawater,F
lorida
,USA
Garay
etal.(20
17b)
Rho
dotorula
kratochvilo
vae
UCDFS
T05
-503
2016
Chy
soperlacareae
(green
lacewing),C
alifornia,
USA
Garay
etal.(20
17b)
Rho
dotorula
aff.p
alud
igena
UCDFS
T81
-84
2016
Opu
ntia
sp.,Ba
hamas
Garay
etal.(20
17b)
Rho
dotorula
paludigena
UCDFS
T09
-163
2016
Leaf
ofDesmod
ium
repens,N
ethe
rlan
dsGaray
etal.(20
17b)
Rho
dotorula
paludigena
UCDFS
T81
-492
2016
Opu
ntia
ficus-indica
cactus,A
rizo
na,U
SAGaray
etal.(20
17b)
Rho
dotorula
paludigena
UCDFS
T82
-646
.220
16Melocactusintortus,P
rickly
Pear
Island
,BritishVirgin
Island
sGaray
etal.(20
17b)
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
399
polyalcohol. Twenty eight years later, a patent was published (Tabuchiand Kayano, 1992) describing production of extracellular esters ofarabitol and mannitol linked to 1–5 molecules of 5-hydroxy-2-decenoicacids by Aureobasidium pullulans strain IFO 7757, Aureobasidium man-sonii strain ATCC 16621, Aureobasidium microstictum strain IFO 32069and a strain of Aureobasidium sp. identified as A-91 or FERM P-10530(Tabuchi and Kayano, 1992). Nagata et al. (1993) screened about 80Aureobasidium sp. strains using a medium containing mannitol as themain carbon source and observed 22 strains producing extracellularheavy oils, which were characterized as 5-hydroxydecenoyl and 3,5-dihydroxydecanoyl arabitol- and mannitol lipids. Kurosawa et al.(1994) confirmed the structures of the heavy oil produced by Aur-eobasidium sp. strain A-2, isolated from the exudate of a tree in the IzuPeninsula (Japan).
In addition to studies of natural production, some strain develop-ment studies have been published. Since the productivity of strain A-2was unstable and decreased with strain storage, an induced mutant,identified as strain A-21M of Aureobasidium sp., was used for furtherstudy.
Studies of liamocin producing fungi have also been connected withtaxonomy and ecology studies. Manitchotpisit et al. (2009) performed amultilocus phylogenetic analysis of 53 fungal isolates, 45 of which werecollected from 15 provinces in Thailand between 2005 and 2006. Thiswork resulted in classification of the fungal isolates of A. pullulans into12 clades. These authors also identified 20 strains capable of secretingheavy oils that would sink to the bottom of the culture. In 2011, thesame group added one more fungal isolate, namely CU 43 (NRRL50380), which was a heavy oil producer as well (Manitchotpisit et al.,2011). They also found that the best producers clustered into clades 8, 9and 11. In 2012, another phylogenetic screening study involving a totalof 56 strains resulted in the creation of two more clades (totaling 14).Four other fungal isolates were identified in this study as heavy oilproducers, confirming liamocin production in 12 of the 14 clades of A.pullulans (Manitchotpisit et al., 2012). Sequencing the BT2 (β-tubulin)locus proved useful in classifying A. pullulans isolates into phylogeneticgroups (Manitchotpisit et al., 2012, 2009). Interestingly, this study in-cluded strains isolated from subpolar environments, whereas the pre-vious screening studies included only strains from tropical environ-ments. In 2014, liamocin production by nine isolates of A. pullulansfrom clades 8, 9 and 11 was examined (Manitchotpisit et al., 2014).Fungal isolates from Clade 11 had the highest liamocin yield. In 2015,A. pullulans strain L3-GPY isolated from a tiger lily, Lilium lancifoliumThunb. in Korea was reported to produce a non-mannitol liamocin,though in low yield (Kim et al., 2015). Strikingly, most liamocin-pro-ducing strains used in these studies (see Table 1) were isolated fromplants or substrates related to plant surface interactions, consistent withthe reputation of A. pullulans as a common epiphyte (Manitchotpisitet al., 2012). These results also showed a strong strain to strain varia-tion in liamocin production.
2.2. Structures of liamocins
The term liamocin was first used by Price et al. (2013) after de-termination of the detailed structure of this new family of compounds.They were previously referred to as “heavy oils” or “extracellular li-pids”. Liamocins consist of a single partially acetylated polyol headgroup, with polyester tails ranging from one to five 3,5-dihydrox-ydecanoic ester groups, which may or may not contain 3-O-acetylationsin the first acyl group (Price et al., 2017) (Fig. 1). Four of the morerecurrent liamocins across the different isolates of A. pullulans are lia-mocins A1, A2, B1 and B2 (Fig. 1) (Leathers et al., 2015; Manitchotpisitet al., 2014, 2011; Price et al., 2013), with [M + Na]+ adduct ions inmatrix-assisted laser desorption/ionization–time-of-flight mass spec-trometry (MALDI-TOFMS) being m/z 763.2, 805.2, 949.4 and 991.4,respectively. Liamocins C1 (m/z 1136) and C2 (m/z 1177.9), consistingof mannitol head groups with pentamers of 3,5-dihydroxydecanoic
ester groups have also been reported in lesser amounts (Bischoff et al.,2015b).
The polyol head group can vary depending on the strain (Bischoffet al., 2015b; Skory et al., 2016), the culture media (Bischoff et al.,2015b; Leathers et al., 2015) and the type of polyol sourced into themedia, but not on the type of sugar sourced into the media (Price et al.,2017). For example, when grown on sucrose as the carbon source,strain CU 43 (NRRL 50380) produces only mannitol-type liamocin (A1,A2, B1, B2 and in less proportion C1 and C2). Strain RSU 12 (NRRL50381) however produces a mixture of mannitol-type liamocin andarabitol-type liamocin when grown in the same medium using sucroseas carbon source (Bischoff et al., 2015b). Growth of strain RSU 6 (NRRL50383) (Bischoff et al., 2015b; Leathers et al., 2015) in productionmedium (PM, described below) (Manitchotpisit et al., 2011) results inproduction of mannitol-type liamocin, whereas growth of the samestrain in a medium high in sea salts (Doshida et al., 1996) results inproduction of mannitol, arabitol and glycerol-type liamocin (Bischoffet al., 2015b; Leathers et al., 2015). Interestingly, strain RSU 12 (NRRL50381) grown in a medium high in sea salts (described below) resultedin a different liamocin profile compared to growth in PM. The stereo-chemistry of the arabitol head group in the resulting liamocin is D-arabitol, rather than L-arabitol (Bischoff et al., 2015b). The commonstereoisomer found in nature is L-(−) (Bischoff et al., 2015b). Growthon D-sorbitol or D-glycerol produced a mixture of liamocins with headgroups primarily containing either mannitol or the substrate polyol(Price et al., 2017). Growth on D-xylitol or D-ribitol produced a mixtureof liamocins containing mannitol, arabitol and substrate head groups.Chiral differences were also observed. Growth on D-threitol producedalmost entirely mannitol-type head groups, whereas growth on L-threitol produced a mixture of mannitol and threitol types. Growth in L-arabitol produced liamocins with a mixture of arabitol and mannitolhead groups, whereas growth on D-arabitol produced liamocins withmostly arabitol head groups (Price et al., 2017). Growth on sugars suchas sucrose, lactose, D-fructose, D-glucose, D-galactose, D-mannose, D-xy-lose and D-arabinose produced only liamocins with mannitol headgroups (Price et al., 2017). These findings suggest that the nature andstereochemistry of the polyols used may play a role in liamocin pro-duction. Sugars and certain polyols may be metabolized and utilized forproduction of mannitol, while other polyols may be partially or com-pletely incorporated directly into liamocins (Price et al., 2017).
The 3,5-dihydroxyacyl ester group was primarily ten carbons inlength, and in some cases the presence of 5-hydroxy-2-decenoic acid (adouble bond between carbons Δ2 and Δ3 instead of the 3-hydroxygroup, Fig. 1) have been reported (Kurosawa et al., 1994; Nagata et al.,1993; Tabuchi and Kayano, 1992), but not confirmed (Price et al.,2013). The hydroxy groups of the polyol head group were not acety-lated, as well as the Δ3 hydroxy groups of subsequent 3,5-dihydrox-ydecanoic ester groups esterified to carbon Δ5 of the first 3,5-dihy-droxydecanoic ester group. The Δ5 hydroxy group of the last 3,5-dihydroxydecanoic ester group was also not acetylated.
Different strains produce different liamocin profiles. For example,strains CU 31, CU 35, CU 39, CU 47 (Manitchotpisit et al., 2011) andstrains in clade 11 (Manitchotpisit et al., 2014) produced a liamocinprofile which had primarily liamocin A2 (m/z 805.2). Strains NRRL Y-2581, NRRL Y-4026 and NRRL 4588 had primarily liamocin B1 (m/z949.4). Strains CU43, CU45 and NRRL Y-12974 had similar amounts ofboth liamocins A2 and B1. Other liamocins were present in smallerproportions across the different fungal isolates. For example, liamocinA1 (m/z 763.2) was present in all strains, while liamocin B2 (m/z991.4) was present in minor proportions in strain CU43, CU45, NRRL Y-2581, NRRL Y-4026, NRRL Y-4588 and NRRL Y-12974 (Manitchotpisitet al., 2011, 2014).
Two analytical approaches have been successful in establishing thestructural features of liamocins. Both have incorporated analysis on thewhole molecule (e.g. mass spectrometry (MS) and nuclear magneticresonance (NMR)), coupled with breakdown analysis of the molecule's
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
400
constituents (e.g. hydrolysis of the polyol, the different 3,5-dihydrox-ydecanoic groups and the acetate groups). The first one, established byTabuchi and Kayano (1992) and used later by Kurosawa et al. (1994)comprised a fractionation step via thin layer chromatography followedby fast atom bombardment mass spectrometry to identify the majorprotonated molecule [M + H]+ in strain A-91, A-2 and A-21M. The finedetails of the structure were elucidated using infrared spectroscopy and1H NMR. Mannitol and arabitol were further confirmed using alkalinehydrolysis followed by paper chromatography and high performanceliquid chromatography (HPLC) using the appropriate standards.Kurosawa et al. (1994) included gas chromatography with flame-ioni-zation detection (GC–FID), gas chromatography–mass spectrometry(GC–MS), 13C NMR, and a further step of preparation of lactones toconfirm the structures of strain A-21M. The second approach followedby Price et al. (2013) also involved native molecule analysis and ana-lysis of the component parts of the molecule. Native molecule analysiscomprised first the identification of the masses of the different liamocincompounds using MALDI-TOFMS and quadrupole/time-of-flight massspectrometry (QTOFMS). These techniques allowed identification of thenumber of 3,5-dihydroxydecanoic ester groups, the type of polyol, andwhether or not the molecule had O-acetylations.
The finer details of the structure including the position of thelinkage of the various 3,5-dihydroxydecanoic ester groups to the polyol,and the position of O-acetyl groups on the polyol, were identified usingmultiple types of NMR analysis (e.g. 1H, 13C, 31P, COSY, HSQC, HMBCand DOSY) (Price et al., 2013). Price et al. (2013) also used dry me-thanolysis in acid media coupled to peracetylation to identify thenumber of free hydroxy groups in the hydrolyzed polar headgroup.Interestingly, they found the presence of 5-hydroxy-2-decenoate lac-tone (massoia lactone) in the methanol soluble fraction and in acidhydrolysis, similar to the lactone reported by Kurosawa et al. (1994). Itis thus possible that Kurosawa created the 5-hydroxy-2-decenoate as anartifact of their analytical procedure, and that strain A-21M might notnecessarily produce it. More research is needed to confirm or discardthis observation. Although liamocin fractionation has been successful, itwas not used towards structure identification, but rather for the de-termination of anti-bacterial activity, as discussed below (Bischoffet al., 2015b).
Exophilins, which are poly(3,5-dihydroxydecanoic ester) groups
also linked via the Δ5 hydroxy group and the carboxyl group withoutany polyol head group, have also been shown to be present in smalleramounts along with liamocins (Price et al., 2013) in a number of iso-lates of A. pullulans. However, these compounds are beyond the scope ofthe present review, and the reader is referred to the following literature(Abdel-Lateff et al., 2009; Chen et al., 1996; Doshida et al., 1996; Wuet al., 2009) for more information on exophilins.
2.3. Growth conditions, substrates and downstream processing
Appropriate media and growth condition are crucial for productionof liamocins, which are secondary metabolites. Liamocins are producedby some Aureobasidium pullulans strains when grown in nitrogen de-prived media in stationary phase. This behavior has been observed inproduction of other fungal glycolipid biosurfactants, such as sophor-olipids (Van Bogaert et al., 2007b) and mannosylerythritol lipids(Morita et al., 2009; Rau et al., 2005). To date, four types of culturemedia have been used to induce production of liamocins (Leatherset al., 2015). For a comparative view of each media, including theconcentrations of each ingredient, see Appendix A. The first one, de-scribed by Manitchotpisit et al. (2009) as production medium (PM), hasbeen extensively used in liamocin research. PM (Leathers et al., 2015) isa relatively simple medium, containing only seven ingredients, in-cluding water and the main carbon source (e.g. sucrose). This mediumhas been used to identify additional liamocin-producing A. pullulansstrains (Manitchotpisit et al., 2011, 2012), to elucidate liamocinstructure (Price et al., 2013), to target the highest liamocin-producingstrains (Manitchotpisit et al., 2014) and the surface tension of liamocins(Manitchotpisit et al., 2011), to identify liamocin antibacterial (Bischoffet al., 2015a; Leathers et al., 2016) and anticancer effects(Manitchotpisit et al., 2014), to test the effect of media composition inliamocin production and diversity (Leathers et al., 2015), and to assessthe effect of different sugars and polyols as carbon sources in the pro-duction of liamocins with different head groups (Price et al., 2017). PMcontains yeast extract and peptone as organic nitrogen sources and noinorganic nitrogen sources. The media is buffered with K2HPO4 andcontains MgSO4·7H2O and NaCl as inorganic salts. The pH is 6.5(Bischoff et al., 2015b; Leathers et al., 2015), and it contains no othermicronutrients or vitamins except those supplied by the yeast extract
Fig. 1. Structures of liamocins and polyol esters of fattyacids.
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
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and the peptone. The medium has been used with 5% w/v of differentcarbon sources. No other carbon source concentration has been pub-lished so far.
Pure sugars, polyols and agricultural wastes have been used ascarbon sources with this media. Interestingly, when wheat straw pre-treated with alkaline hydrogen peroxide was used as the carbon source,the resulting liamocins were free of melanin contamination previouslyobserved in cultures grown with sucrose (Leathers et al., 2016). Al-though liamocin yield was low using pretreated wheat straw with al-kaline hydrogen peroxide as the carbon source, addition of hydrolyticenzymes prior to fermentation increased production, to a point where itbecame comparable to sucrose (Leathers et al., 2016). The liamocinprofile also changed, giving place to higher abundance of non-acety-lated liamocins A1 and B1, which have the highest biological activityagainst Streptococcus sp. (Leathers et al., 2016).
The initial carbon to nitrogen (C:N) ratio is an indicator that hasbeen used in single cell oil research to assess lipogenesis potential inoleaginous microorganisms (Garay et al., 2014). For yeast, it has beensuggested that ideal lipid production occurs in media having initial C:Nratios between 30 and 80, depending on the yeast species and thecarbon source (Moreton, 1988; Sitepu et al., 2014). Initial C:N ratiosbelow or above this range may result in poor single cell oil production.Interestingly, initial C:N ratios that are optimal for production otherEFGB such as sophorolipids display a wider range than that suggestedfor single cell oil production. For comparison, initial C:N ratios reportedfor sophorolipid production are 48 (Davila et al., 1992), 107 (Pekinet al., 2005), 140 (Rau et al., 2001), and 529 (Daniel et al., 1998). Inthese experiments, 320 g/L,> 400 g/L, 300 and 422 g/L were ob-tained, with conversions of 0.65, 0.60, 0.68 and 0.84 g sophorolipid pergram of carbon source, respectively (Table 2). Unlike polyol lipids, SLproduction is achieved by providing both a sugar and a hydrophiliccarbon source (Van Bogaert et al., 2007b). Interestingly, these experi-ments were performed in fed batch bioreactors where an excess carbonsource was added after the fermentation reached stationary phase, forcontinued conversion of the carbon source into product by the maturecells. Assuming that peptone has a carbon fraction of 0.31 (Rajasekaranet al., 2011) and a nitrogen fraction of 0.13, and assuming that thecarbon fraction in yeast extract is 0.4 (Holwerda et al., 2012), and thenitrogen fraction is 0.11, then the C:N ratio of this medium when using5% glucose (or sucrose) would be 166. The highest yields using thismedium have been 10.4 and 10.6 g/L by strains CU 31 and CU 42 re-spectively (see Table 2), from 50 g/L sucrose grown in 50 mL culturesin 250 mL shake flasks incubated for 7 days at 28 °C (Leathers et al.,2016). These values translate into productivities of 0.208 and 0.212 gliamocin per gram of sucrose. Strains RSU 21 and RSU 29 are also highliamocin producers (reported yields are 8.4 and 8.6 g/L respectively)(Manitchotpisit et al., 2014) classified in Clade 11, that also producehigh amounts of pullulan: 17 and 9.4 g/L respectively (Manitchotpisitet al., 2014).
The second media is also a relatively simple media, containing eightingredients including water and the carbon source. It was first docu-mented in 1992 (Tabuchi and Kayano, 1992) with slight variations insubsequent publications (Kurosawa et al., 1994; Nagata et al., 1993).This medium has a mixture of organic and inorganic nitrogen sources(see Appendix A). The pH is 5.5, but adjustment to 6.5 increased lia-mocin production (Tabuchi and Kayano, 1992). Kurosawa et al. (1994)reported a rough yield estimate of about 35 g/L produced by Aur-eobasidium sp. strain A-21M after combining the medium of twenty500 mL flasks containing 50 mL of media each. However, six otheryeast strains, namely, A. pullulans CU 43 (NRRL 50380), RSU 12 (NRRL50381), RSU 32 (NRRL 50382), RSU 6 (NRRL 50383), RSU 29 (NRRL50384) and NBRC 10874 (NI10102) were tested using the same media,and the highest yield was 4.8 g/L (Leathers et al., 2015), achieved bystrain RSU 29 (NRRL 50384), just 14% of that reported by Kurosawa.The main carbon source used with this medium was glucose at a 12%w/v concentration. However, other minor carbon sources used with this
medium in screening steps include corn steep liquor, succinic acid andammonium succinate. The C:N ratio of this medium is 112. Interest-ingly, Kurosawa et al. (1994) reported that addition of the neutralizingagent CaCO3 increases production of poly-(β-L-malic acid) (PMA) anddecreases liamocin secretion. The same phenomenon was observed byManitchotpisit et al. (2012). They concluded that although culture pH isstabilized by the addition of CaCO3, it is not clear that pH is the solefactor influencing the ratio of PMA/liamocin production(Manitchotpisit et al., 2012). Lee et al. (1999) found that 13C fromCa13CO3 was indeed present in PMA synthesized by the slime moldPhysarum polycephalum, suggesting that carbon from CaCO3 is used bythe slime mold as carbon source to synthesize PMA. More research isneeded to confirm this attribute in A. pullulans.
The third medium was first used by Doshida et al. (Doshida et al.,1996) originally for production of exophilin A, and is the simplest of thefour media used for liamocin production, containing only L-aspar-agine·H2O, sea salts, K2HPO4, water and a carbon source. The pH is 7.0.The only carbon source and only concentration thus far reported forliamocin production is 1% glucose, being 5 and 12 times less than theprevious two media. Interestingly, strain NRRL 50384 produced 1.4 g/Lliamocins from 10 g/L glucose, equivalent to a productivity of 0.14 gliamocins per gram of glucose. This value ranks among the highestproductivities reported so far for liamocin production (Table 2). TheC:N ratio is 37.5.
The fourth medium is the most complex (Leathers et al., 2015; Wanget al., 2014) and contains inorganic and organic nitrogen sources, and acocktail of micronutrients (See Appendix A). The pH is 6.0. Thismedium has the highest C:N ratio: 496.
Liamocin productivity varied depending on the type of carbonsource used. For example A. pullulans strain CU 43 (NRRL 50380)grown using the same basal media (e.g. PM media) and growth con-ditions, produced the following yields from the following carbon sub-strates (in decreasing order): sucrose, 4.4 g/L; oat spelt xylan, 3.1 g/L;D-glucose, 2.5 g/L, pretreated wheat straw, 1.6 g/L; D-mannitol, 0.6 g/Land lactose, 0.2 g/L (Price et al., 2017). The carbon substrates weredosed at 50 g/L in all cases (Leathers et al., 2016; Price et al., 2017).Interestingly, when fed pretreated wheat straw as the carbon source,the resulting liamocins generally were less acetylated than when grownon sucrose: the ratios of acetylated tail group species A2 and B2 wererelatively lower in liamocins produced on pretreated wheat straw thanon sucrose (Leathers et al., 2016).
Few comparisons have been conducted to assess the optimal con-ditions for liamocin production for parameters such as pH, temperature,C:N ratio, culture to volume ratio/aeration, incubation time and in-oculum ratio. In terms of pH, the starting pH used so far has rangedfrom pH 5.5 to pH 7, and biological acidification of the media (probablydue to the yeast secreting organic acids) has been reported to achievevalues in the range of 5.5 to as low as pH 3 (Manitchotpisit et al., 2012;Nagata et al., 1993). There is not a clear picture on whether higheryields can be obtained at initial media pH values closer to 7 or closer to5.5. A narrow temperature range (25–30 °C) has been used for liamocinproduction, and the two most frequent temperature values have been25 and 28 °C (Kurosawa et al., 1994; Leathers et al., 2015;Manitchotpisit et al., 2011). To our knowledge no study targetingtemperature optimization has been published yet. However, the highestliamocin yield has been obtained at 25 °C (Kurosawa et al., 1994).
Most published studies have used culture to volume ratios rangingbetween 1:3 and 1:10. For example, Manitchotpisit et al. (2011) con-ducted experiments using 80 mL cultures in 250 mL flasks (for a cultureto volume ration of 1:3.2) with yields as high as 6.0 g/L. The highestyields have been reported compiling twenty 50 mL cultures each in500 mL flasks, equivalent to a culture to volume ratio of 1:10(Kurosawa et al., 1994). These results suggest that aeration is a keyfactor for liamocin production. While the present work was under re-view, a central composite face-centered study using a stirred tankbioreactor identified that liamocin yield (expressed as the maximum
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
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Table2
Polyol
lipid
substrateco
nversion
andmultipleco
-produ
ctprod
uctivity
repo
rted
bytopprod
ucingAscom
ycetou
san
dBa
sidiom
ycetou
sspecies.
Speciesna
me
Strain
Type
ofEF
GB
Totalcarbon
supp
lied(g/L
)Culture
metho
dEF
GB
yield(g/L
)Yield
ofothe
rco
-produ
cts(g/
L)
EFGBco
nversion
(gEFG
B/g
carbon
source)
Totalco
nversion
(gtotalproducts/g c
arbon
source)
EFGBprod
uctivity
(mg E
FGB/[g c
arbon
source×
day])
Totalprod
uctivity
(mg T
otal
products/[g
carbon
source×
day])
Referen
ce
Aureoba
sidium
sp.
A-21M
LC12
0gluc
ose
Flask
35N/A
0.29
0.29
41.7
41.7
Kurosaw
aet
al.
(199
4)A.p
ullulans
NRRLY-
1297
LC50
gluc
ose
Flask
1.9
PMA:6
.7Pu
llulan:
190.04
0.55
5.4
78.9
Man
itch
otpisitet
al.
(201
2)A.p
ullulans
RSU
9LC
50gluc
ose
Flask
7.0
Pullu
lan:
140.14
0.42
2060
Man
itch
otpisitet
al.
(201
2)A.p
ullulans
RSU
21LC
50gluc
ose
Flask
8.6
Pullu
lan:
170.17
0.51
24.6
73.1
Man
itch
otpisitet
al.
(201
2)A.p
ullulans
RSU
29LC
50gluc
ose
Flask
8.4
Pullu
lan:
940.17
0.36
2450
.9Man
itch
otpisitet
al.
(201
2)A.p
ullulans
RSU
29LC
10gluc
ose
Flask
1.4
N/A
0.14
0.14
2020
Leathe
rset
al.(20
15)
A.p
ullulans
CU
31LC
50gluc
ose
Flask
10.4
N/A
0.21
0.21
29.7
30.3
Leathe
rset
al.(20
16)
A.p
ullulans
CU
42LC
50gluc
ose
Flask
10.6
N/A
0.21
0.21
29.7
30.3
Leathe
rset
al.(20
16)
A.p
ullulans
RSU
9LC
50whe
atstraw
with
enzymes
Flask
6.5
N/A
0.13
0.13
18.6
18.6
Leathe
rset
al.(20
16)
R.a
ff.p
alud
igena
UCDFS
T81
-84
PEFA
50gluc
ose
Flask
12.4
N/A
0.25
0.25
35.4
35.4
Garay
etal.(20
17b)
R.p
alud
igena
UCDFS
T81
–492
PEFA
50gluc
ose
Flask
11.7
N/A
0.23
0.23
33.4
33.4
Garay
etal.(20
17b)
R.b
abjevae
UCDFS
T04
-83
0PE
FA50
gluc
ose
Flask
8.5
N/A
0.17
0.17
24.3
24.3
Garay
etal.(20
17b)
R.a
ff.p
alud
igena
UCDFS
T81
-84
PEFA
50gluc
ose
Bioreactor
(Batch
)12
.3TG
:3.2
0.22
0.27
3544
Garay
etal.(20
17a)
R.a
ff.p
alud
igena
UCDFS
T81
-84
PEFA
100gluc
ose
Bioreactor
(Batch
)14
.5TG
:6.3
0.16
0.24
20.7
29.6
Garay
etal.(20
17a)
R.a
ff.p
alud
igena
UCDFS
T81
-84
PEFA
150gluc
ose
Bioreactor
(Fed
Batch)
20.9
TG:8
.80.16
0.24
19.9
28.2
Garay
etal.(20
17a)
R.a
ff.p
alud
igena
UCDFS
T81
-84
PEFA
150sucrose
Bioreactor
(Fed
Batch)
31.2
N/A
0.21
0.21
29.7
29.7
Unp
ublishe
dda
ta
R.a
ff.p
alud
igena
UCDFS
T81
-84
PEFA
100glycerol
Bioreactor
(Batch
)18
.7N/A
0.19
0.19
26.7
26.7
Unp
ublishe
dda
ta
R.p
alud
igena
UCDFS
T81
-49
2PE
FA10
0gluc
ose
Bioreactor
(Batch
)17
.5N/A
0.17
0.17
25.0
25.0
Unp
ublishe
dda
ta
Starmerella
bombicola
ATC
C22
214
SL20
0gluc
ose,
450
CornOil20
Hon
eyBioreactor
(Fed
Batch)
>40
0N/A
0.60
0.60
33.3
33.3
Pekinet
al.(20
05)
Starmerella
bombicola
ATC
C22
214
SL10
0lactose,
1gluc
ose,
2ga
lactose,
400rape
seed
oil
Bioreactor
(Fed
Batch)
422
N/A
0.84
0.84
49.1
49.1
Dan
ielet
al.(19
98)
Starmerella
bombicola
ATC
C22
214
SL10
0gluc
ose,
3.5mL/
hrape
seed
ethy
lesters
Bioreactor
(Fed
Batch)
320
N/A
0.65
0.65
81.3
81.3
Dav
ilaet
al.(19
92)
Starmerella
bombicola
ATC
C22
214
SL30
0gluc
ose,
140
rape
seed
oil
Bioreactor
(Fed
Batch)
300
N/A
0.68
0.68
130.6
130.6
Rau
etal.(20
01)
Abb
reviations:L
C:lia
moc
ins;
PEFA
:polyo
lesters
offattyacids;
EFGB:
Extracellularfung
alglycolipid
biosurfactan
ts;TG
:triacylglycerols;
PMA:p
oly-(β- L-m
alic
acid).
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
403
biosurfactant tensoactivity increases with increased aeration, obtainingthe highest value at 1.1 L/min (Pereira Brumano et al., 2017). It hasbeen a common practice to incubate cultures of A. pullulans for liamocinproduction for 7 days. However, a shake flask time course experimentthat lasted for 14 days showed a decrease in liamocin yield after10 days (Leathers et al., 2016). Inoculum ratios ranging from 8 to 10%have been used for liamocin production (Kurosawa et al., 1994; Nagataet al., 1993).
Different strategies for liamocin recovery in shake flask level havebeen reported. The easiest approach, which results in the highest yields,involves solvent extraction of the full culture in culture-to-solvent ratiosvarying from 1:1 to 1:2, followed by phase separation and evaporationof the solvent layer to dryness (Method 1) (Leathers et al., 2016, 2015;Price et al., 2017). Another strategy involves a first step of centrifuga-tion to separate the spent media, followed by further separation of thecell layer to extract the remaining liamocins using a solvent, or ex-tracting the cell layer and liamocins together with a solvent, finishingwith evaporation, either under a stream of air or under reduced pres-sure (Method 2). The latter method was used to report simultaneousyields of pullulan, PMA and liamocins (Manitchotpisit et al., 2012).However, since pullulan and PMA are water soluble polymers, theformer method plus further separation of the water phase followed bytreatment with ethanol could be a strategy to recover three valuable co-products from A. pullulans cultures. The most common solvent used toextract liamocins is methyl ethyl ketone (butan-2-one) (Leathers et al.,2015; Manitchotpisit et al., 2011).
Other solvents notably ethyl acetate have been used as well(Tabuchi and Kayano, 1992). Melanin pigments have been reported tocoextract with liamocins from cultures containing sucrose as carbonsource. Interestingly, it was reported that the use of pretreated wheatstraw with alkaline hydrogen peroxide as carbon source resulted inliamocins devoid of melanin pigments (Leathers et al., 2016). In onestudy, a red pigment co-extracted in the methyl ethyl ketone fractionfrom A. pullulans strain RSU 12 (NRRL 50381) and was identified tohave an [M+ Na]+ ion m/z 476 based on MALDI-TOF analysis(Bischoff et al., 2015b). Downstream processing for liamocins appearseasier and less cumbersome compared to other EFGB, such as SL andMEL lipids, because these two require hydrophobic carbon sources forproduction, and removal of the remaining unused hydrophobic sourceusually involves an additional purification step (Morita et al., 2009;Van Bogaert et al., 2011). In addition to a more complicated down-stream processing pathway, addition of hydrophobic substrates forother types of EFGB production bears a substantial cost in the economicmodel, as has been shown for the case of SL (Ashby et al., 2013).However, high glycolipid production in part compensates for the cost ofinput materials. Thus, production of EFGB from hydrophilic substratesis a beneficial feature that has been explored in the case MEL (Moritaet al., 2015).
2.4. Physiological role of liamocins
There have been no experiments conducted to demonstrate thephysiological role and function of liamocins for A. pullulans. However, ithas been suggested that EFGB secretion play a role in conditioning leafsurfaces making them a suitable environment for microbial growth byenhancing the permeability of the outer epidermis wall (Ruinen, 1963).Further experiments (Bunster et al., 1989; Schreiber et al., 2005) areconsistent with this theory. The observation that the majority of thestrains that secrete liamocins were isolated from the phyllosphere orsome sort of plant surface interaction source is also consistent with thistheory. The fact that liamocins display selective antibacterial effectcould also constitute a competitive advantage for A. pullulans. Lipasesecretion by A. pullulans has been well documented (Federici, 1982;Leathers et al., 2013; Liu et al., 2008). Lipase secretion, lipase activityand liamocin production by A. pullulans all peak around 5 to 6 dayscultivation (Leathers et al., 2013; Manitchotpisit et al., 2011), which
supports the hypothesis that liamocins are an extracellular form ofcarbon storage. Since liamocins contain polyester tails ranging from oneto five 3,5-dihydroxydecanoic ester groups, a controlled and targetedrelease of either 3,5-dihydroxydecanoic pentamers, tetramers, trimers(e.g. exophilin A, molecules that display antibacterial activity) (Doshidaet al., 1996), could help the fungus ensure carbon availability that noother competing microbes could use during periods of carbon stress. Asimilar function for SL has been proposed elsewhere (Van Bogaert et al.,2007b). In this case, Starmerella bombicola, the highest SL yeast pro-ducer thus far reported has been successfully grown in SL as sole carbonsource (Garcıa-Ochoa and Casas de Pedro, 1997). It remains to be seenwhether A. pullulans can grow in liamocins as the sole carbon source aswell.
Literature in the late 1950's suggested that A. pullulans played a roleas plant pathogen causing “diseases of legumes and flax” and “a scald ofgrape leaves” (Cooke, 1959). A report in the early 1990's concluded thatthe literature reports for A. pullulans involvement in pathogenesis ofplants had pros and cons (Deshpande et al., 1992). More recent studieshave focused less on the pathogenic side and more on the effect ofcertain A. pullulans strains as biocontrol against plant pathogens fordifferent industrially important crops, like apples (Schena et al., 1999)and peaches (di Francesco et al., 2015). These are different strains fromthe liamocin-producing strains described in the present work. Furtherresearch is needed to prove whether a connection between liamocinproduction and A. pullulans biocontrol behavior exists.
2.5. Biosynthesis of liamocins
The liamocin full biosynthetic pathway is unknown. However, ahypothetical biosynthetic pathway was proposed for production of amannitol liamocins with a single non-acetylated 3,5-dihydrox-ydecanoate group by A. pullulans strain 5P (Li et al., 2015) (Fig. 2). Inthis case, malonyl-CoA derived from acetyl-CoA is condensed by 3-ke-toacyl synthase (KS), a condensing subunit present within a polyketidesynthase (PKS) multicomplex, to form a C-4 carbon moiety. The nascentmolecule undergoes a fully reductive cycle involving reduction via 3-ketoacyl-ACP reductase (KR), dehydration via dehydratase (DH) and alast reductive step via enoyl reductase (ER). Another malonyl-CoA iscondensed to the molecule to have now a six carbon moiety that willundergo a fully reductive cycle. From this point on, two more con-densations via KS followed by subsequent reductions with KR will yielda single 3,5-dihydroxydecanoate group still bound to the acyl carrierprotein (ACP) of PKS. This pathway suggested by Li et al. (Li et al.,2015) requires 3-hydroxydecanoyl-ACP:CoA transacylase (PhaG) en-zyme, encoded by a phaG gene to release the 3,5-dihydroxydecanoategroup from ACP as a CoA derivative (see Fig. 2), before the in-corporation of mannitol. However, a proof of this step remains to bedone in liamocin-producing strains.
The genome of A. pullulans has been sequenced, and genes for type IPKS genes were found in four different sub-varieties of A. pullulans(Gostinčar et al., 2014). Genes for phaG were also found in a genomesequenced from strain AY4 (Li et al., 2015). However, cloning andcharacterization of these enzymes in liamocin-producing strains areneeded to confirm the latter hypothesis.
Cultures of A. pullulans strain NRRL 50380 were grown on uni-versally isotopically-labeled [U-13C] glucose, and liamocins were ana-lyzed using MALDI-TOFMS (Price et al., 2013). The 13C-isotope fromthe labeled glucose was found in the mannitol part of the molecule, butnot in the 3,5-dihydroxydecanoate group (Price et al., 2013). This in-formation suggests that glucose is metabolized into mannitol for lia-mocin production, and that the 3,5-dihydroxydecanoic ester group israther complex.
Mannitol has been shown to be derived from glucose (Skory, 2016)and two important enzymes have been shown to play a key role on theproduction of mannitol head groups for liamocin biosynthesis in A.pullulans (Fig. 2): mannitol-1-phosphate dehydrogenase (MDP1) and
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
404
mannitol dehydrogenase (MDH2) (Skory, 2016). Genetically modifiedknock-out strains of A. pullulans were created targeting the removal ofcombinations of four genes from wild type strain A. pullullans RSU 6(NRRL 50383) and RSU 29 (NRRL 50384), resulting in either the pro-duction of arabitol-liamocins, melanin free liamocins and the discoveryof synthesis of massoia lactone (Skory et al., 2016). The genes werempd, two types of mdh, namely mdh1, a gene encoding a putativemannitol dehydrogenase that is likely a cytochrome enzyme apparentlylocated in the mitochondria, responsible for converting mannitol to
fructose, mdh2 a gene encoding another mannitol dehydrogenase in A.pullulans, and pks4, a gene encoding a functional component of PKSinvolved in melanin production, a pigment that is co-extracted alongwith liamocins during solvent extraction (Skory et al., 2016).
Further research is needed to understand the esterification of thehydroxylated fatty acid to the sugar alcohol that occurs in PEFAsynthesis, as well as the order of these esterifications. Li et al. (2015)suggested that the last step in PEFA biosynthesis in Aureobasidium iscarried out by an esterase, but failed to provide further information as
Fig. 2. Suggested biosynthetic pathway of amannitol-type liamocin with a single 3,5-dihy-droxydecanoic ester group, adapted from Li et al.(2015) and Skory et al. (2016). MDP1: mannitol-1-phosphate dehydrogenase; MPP: mannitol;MDH2: Mannitol dehydrogenase (Skory et al.,2016); KS: 3-ketoacyl synthase subunit of type Ipolyketide synthase (PKS); KR: 3-ketoacyl-ACPreductase; DH: dehydratase subunit of type I PKS;ER: enoyl reductase subunit of type I PKS; PhaG:3-hydroxydecanoyl-ACP:CoA transacylase (Liet al., 2015).
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
405
to where the esterase is located and its mechanism of action. More workis also required to elucidate the liamocin mechanism of secretion.
2.6. Properties, uses and applications of liamocins
Liamocins have many commercially relevant properties. They havebeen described as oils that are heavier than water that appear as an oilylayer beneath the cell layer after centrifugation. The color of liamocinsrange between bright yellow and various shades of greens(Manitchotpisit et al., 2011), to light or even dark brown (Kurosawaet al., 1994) depending on the strain. Thirteen strains have been re-ported to display fluorescence (Manitchotpisit et al., 2011), not asso-ciated with the presence of riboflavin nor aromaticity (Manitchotpisitet al., 2011). Surface tensions (Table 3) for liamocins have been re-ported to range between 27 mN/m (Manitchotpisit et al., 2011) and31.5 mN/m (Kim et al., 2015). The surface tension of pure water is72–73 mN/m at 20–25 °C (Pallas and Harrison, 1990). These values forreduction of surface tension are in the range of commercial surfactants(Table 3).
Liamocins have also been reported to display an antiproliferativeeffect against certain cancer cell lines (Isoda et al., 1997; Isoda andNakahara, 1997; Manitchotpisit et al., 2014). A study in 1997 (Isodaet al., 1997) following the method of Kurosawa et al. (1994) showedthat liamocins induced cell differentiation into granulocytes instead ofcell growth in the human promyelocytic leukemia cell line HL60. Asimilar behavior occurred when human myelogenous leukemia cell lineK562 and the human basophilic leukemia cell line KU 812 were ex-posed to liamocins, triggering their differentiation into megakaryocytesand granulocytes, respectively (Isoda and Nakahara, 1997). The samegroup of researchers also showed that liamocins inhibited growth ofhuman lung cancer cell line A549, causing a significant decrease in theactivity of intracellular protein kinase C (Isoda et al., 1997). In anotherstudy, liamocins from strains CU43 and NRRL Y-12974 had differenteffects on specific mammalian cell types (Manitchotpisit et al., 2011).Liamocins from strain NRRL Y-12974 inhibited non-cancerous Africangreen monkey kidney cells (control) and cancerous small cell lungcancer cells in similar levels, while almost half the dose was required toinhibit oral cavity cancer cells. In the other hand, liamocins from strainCU43 (NRRL 50380) had no cytotoxicity against the aforementionednon-cancerous cell line, but effectively inhibited the two cancer celllines described above. The two main liamocin molecules present in bothstrains are A2 (m/z 805.5), and B1 (m/z 949.6), suggesting that thedifferences in the cytotoxicity cannot be simply attributed to the mostabundant liamocin produced by the strains. In another study publishedin 2014 (Manitchotpisit et al., 2014), liamocins without further frac-tionation from nine A. pullulans strains were tested for inhibition of twohuman breast cancer cell lines (T47D and SK-BR3) and a human cer-vical cell line (HeLa) (Manitchotpisit et al., 2014). Liamocins fromstrains RSU 20, RSU 17 and RSU 19, RSU 9 and RSU 21 inhibited allthree cancer cell lines, without showing cytotoxicity to a control non-cancerous African green monkey kidney epithelial cell line (Vero)(Manitchotpisit et al., 2014). The half maximal inhibitory concentration(IC50) values exhibited by liamocins ranged from 26.0–83.6 μg lia-mocin/mL whereas IC50 values from other prospective anticanceragents ranged from 25 to 35 μg/mL (Manitchotpisit et al., 2014). Theanticancer effect of individual liamocin molecules has not yet beenverified, though liamocin B1 was present in high proportions in most ofthe mixtures displaying anticancer effect across the latter two studies(Manitchotpisit et al., 2011, 2014).
EFGB are known to display antimicrobial activity. For example, SLhave antimicrobial activity against certain Candida and Pichia yeastspecies, as well as some Gram-positive bacteria (Ito et al., 1980; Langet al., 1989; Van Bogaert et al., 2007b), and MEL have strong anti-microbial activity against Gram-positive bacteria, and some activityagainst gram-negative bacteria (Morita et al., 2009). Liamocins havealso been shown to be powerful selective antimicrobial molecules. TheTa
ble3
Surfacetensionva
lues
fordifferen
textracellularfung
alglycolipid
biosurfactan
ts(EFG
B).T
hesurfacetensionforpu
rewater
is72
–73mN/m
at20
–25°C
(Pallasan
dHarrison,
1990
).
Species&
strain
EFGB
Minim
umsurfacetension(m
N/m
)Metho
dReferen
ce
Aureoba
sidium
pullu
lans
CU
43(N
RRL50
380)
Liam
ocin
(Man
nitolhe
adgrou
p)27
(tem
perature
notrepo
rted
)Pe
ndan
tdrop
metho
dMan
itch
otpisitet
al.(20
11)
Aureoba
sidium
pullu
lans
L3-G
PYLiam
ocin
(Glycerolhe
adgrou
p)31
.5(tem
perature
notrepo
rted
)Ring/
platemetho
dKim
etal.(20
15)
Rho
dotorula
babjevae
UCDFS
T04
-877
PEFA
30.4
(25°C)
Wilh
elmyplatemetho
dXu(201
7)Rho
dotorula
aff.p
alud
igenaUCDFS
T81
-84
PEFA
33.3
(25°C)
Wilh
elmyplatemetho
dXu(201
7)Starmerella
bombicola
ATC
C22
214
Soph
orolipid
37(tem
perature
notrepo
rted
)Wilh
elmyplatemetho
dSo
laim
anet
al.(20
04)
Wickerham
iella
domercqiaeY2A
Soph
orolipid
37.3
(tem
perature
notrepo
rted
)DeNou
ryring
metho
dChe
net
al.(20
06)
Pseudo
zymaan
tarcticaT-34
(previou
slykn
ownas
Can
dida
antarctica)
Man
nosylerythritollip
id28
(25°C)
Wilh
elmyplatemetho
dKitam
otoet
al.(19
93)
Van
rija
humicola9-6(previou
slykn
ownas
Cryptococcushu
micola)
Cellobioselipids
37(23°C)
DeNou
ryring
metho
dPu
chko
vet
al.(20
02)
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
406
Table4
Antibacterial
activities
oflia
moc
inssecreted
bydifferen
tstrains
ofAureoba
sidium
pullu
lans.T
hean
tiba
cteriala
ctivitywas
measuredusingthebrothmicrodilution
metho
d(Bisch
offet
al.,20
15a,
2015
b;Man
itch
otpisite
tal.,
2014
)an
dtheva
lues
are
expressedin
minim
uminhibitory
conc
entration(M
IC)va
lues
(μglia
moc
in/m
L).
Speciesna
mean
dstrain
ID.
A.p
ullulans
RSU
9(N
RRL
6203
1)
A.p
ullulans
RSU
13(N
RRL
6203
4)
A.p
ullulans
RSU
17(N
RRL
6203
8)
A.p
ullulans
RSU
18(N
RRL
6203
9)
A.p
ullulans
RSU
19(N
RRL
6204
0)
A.p
ullulans
RSU
20(N
RRL
6204
1)
A.p
ullulans
RSU
21(N
RRL
6204
2)
A.p
ullulans
RSU
29(N
RRL
5038
4)
A.p
ullulans
RSU
32(N
RRL
5038
2)
A.p
ullulans
CU
43(N
RRL
5038
0)
Referen
ce
Lactobacillus
ferm
entum
0315
-1>
1250
>12
50>
1250
>12
50>
1250
>12
50ND
>12
50>
1250
>12
50Bischo
ffet
al.(20
15a,
2015
b);M
anitch
otpisit
etal.(20
14)
Enterococcus
faecalis
ATC
C29
213
312
625
312
312
312
312
ND
312
156
312
Bischo
ffet
al.(20
15a,
2015
b);M
anitch
otpisit
etal.(20
14)
Stap
hylococcus
aureus
ATC
C29
213
>12
50>
1250
>12
50>
1250
>12
50>
1250
>61
2>
1250
>12
50>
1250
Bischo
ffet
al.(20
15a,
2015
b);M
anitch
otpisit
etal.(20
14)
EscherichiacoliATC
C25
922
>12
50>
1250
>12
50>
1250
>12
50>
1250
ND
>12
50>
1250
>12
50Bischo
ffet
al.(20
15a,
2015
b);M
anitch
otpisit
etal.(20
14)
Pseudo
mon
asaerugino
saATC
C27
853
>12
50>
1250
>12
50>
1250
>12
50>
1250
ND
>12
50>
1250
>12
50Bischo
ffet
al.(20
15a,
2015
b);M
anitch
otpisit
etal.(20
14)
Streptococcusagalactia
eNRRLB-18
1539
7878
3939
39NA
NA
NA
20Bischo
ffet
al.(20
15a,
2015
b)Streptococcusub
eris
NA
NA
NA
NA
NA
NA
NA
NA
NA
78Bischo
ffet
al.(20
15a,
2015
b)Streptococcusmutan
sATC
C25
175
NA
NA
NA
NA
NA
NA
NA
NA
NA
78Bischo
ffet
al.(20
15a)
Streptococcusmitis
NRRLB-14
574
NA
NA
NA
NA
NA
NA
NA
NA
NA
20Bischo
ffet
al.(20
15a)
Streptococcus
infantariusNRRL
B-41
208
NA
NA
NA
NA
NA
NA
NA
NA
NA
78Bischo
ffet
al.(20
15a)
Streptococcussalivarius
NRRLB-37
14NA
NA
NA
NA
NA
NA
NA
NA
NA
≤10
Bischo
ffet
al.(20
15a)
Streptococcussobrinus
NRRLB-44
68NA
NA
NA
NA
NA
NA
NA
NA
NA
>12
50Bischo
ffet
al.(20
15a)
Bacillu
ssubtilisMW10
NA
NA
NA
NA
NA
NA
NA
NA
NA
640
Bischo
ffet
al.(20
15b)
StreptococcussuisATC
C43
765
NA
NA
NA
NA
NA
NA
NA
NA
NA
≤10
Bischo
ffet
al.(20
15b)
Abb
reviations:N
D:N
otde
term
ined
;NA:N
otav
ailable.
L.A. Garay et al. Biotechnology Advances 36 (2018) 397–414
407
antibacterial effect of liamocins has been tested qualitatively via theagar diffusion method and quantitatively via the broth microdilutionmethod against several types of microorganisms. The quantitative re-sults in terms of minimum inhibitory concentration (MIC), in μg lia-mocin/mL, reported for strain A. pullulans CU 43 (NRRL 50380) andnine additional strains are depicted in Table 4 (Bischoff et al., 2015a,2015b; Manitchotpisit et al., 2014). These results show that the anti-bacterial activity is specific towards Streptococcus genus and it appearsto be a general property of Aureobasidium's liamocins. Furthermore, theantimicrobial activity is retained even when liamocins are subjected toheat (e.g. autoclaving) (Bischoff et al., 2015b). Fluorescence assaysshowed that liamocins generate a rapid loss of membrane integrity forStreptococcus agalactiae (Bischoff et al., 2015a), reducing viable bac-terial densities 1.4 log(CFU/mL) within 1 h at a concentration as low as250 μg/mL (Bischoff et al., 2015b). Treatment of S. uberis with156 μg liamocin/mL also reduces viability over 3 log(CFU/mL) in onehour (Bischoff et al., 2015b).
Until 2014, all the experiments testing liamocin antimicrobial andanticancer effects were conducted using crude liamocin mixtures(Manitchotpisit et al., 2014). In 2015, a patent was published de-scribing the liamocin fractionation of A. pullulans strain CU 43 (NRRL50380) into fourteen fractions using a surveyor reverse phase HPLCsystem equipped with a photodiode array detector (Bischoff et al.,2015b). Fractions seven through thirteen successfully separated in-dividual liamocins A1, A2, B1 and B2. Minor fractions C1 and C2 co-eluted in fraction 14. Each of these fractions was tested for antibacterialactivity against S. agalactiae qualitatively via the disc diffusion assayand quantitatively via the broth dilution method (Bischoff et al., 2015a,2015b). Fraction 12 containing mannitol-liamocin B1 (m/z 949.4) hadthe lowest MIC of 16 μg/mL, followed by fraction 14 with the combi-nation of mannitol-liamocins C1/C2 with 32 μg/mL, and a tie betweenmannitol-liamocins A1 and A2 with 64 μg/mL (Bischoff et al., 2015b).Interestingly, liamocin B1 (m/z 949.4) had the highest antimicrobialeffect and was the most abundant liamocin in the mixtures that alsodisplayed the most effective anticancer activity, as mentioned above.
Liamocins with other types of polyol head groups are also activeagainst Streptococcus spp. (Price et al., 2017). Antibacterial effectagainst S. agalactiae of liamocins containing D-arabitol head groups anda mixture of D-xylitol:D-arabitol (1.67:1) head groups were testedagainst mannitol-liamocins to compare the effect of liamocin headgroup in the antibacterial effect. MIC were higher for D-arabitol and themixture of D-xylitol:D-arabitol head groups suggesting that the anti-bacterial activity of non-mannitol liamocins is weaker than mannitol-liamocins (Bischoff et al., 2015b). All these observations mean thatliamocins with various polyol head groups retain antibacterial specifi-city towards Streptococcus sp. Interestingly, the number of 3,5-dihy-droxydecanoic ester groups appeared to have had an effect in the killingbehavior of arabitol liamocins, since D-arabitol liamocin tetramers de-monstrated better killing activity than D-arabitol trimers (Bischoff et al.,2015b), and dimeric and monomeric forms are inactive against S.agalactiae (Price et al., 2017). Exophilins and halymecins are anti-bacterial molecules secreted by Fusarium spp. and Simplicillium spp.Although they resemble liamocins structurally, they differ in lacking apolyol head group and thus they display a different spectrum of activity(Price et al., 2017).
Liamocin's narrow spectrum of antibacterial activity may be ad-vantageous over broad-spectrum agents, especially in prophylactic ap-plications to target streptococcal pathogens such as S. agalactiae and S.uberis, which have been shown to cause mastitis in dairy cattle(Leathers et al., 2016). Furthermore, liamocins could avoid disruptionof the beneficial normal microbiome or selection for antibiotic re-sistance in commensal bacteria (Bischoff et al., 2015a). Liamocin-con-taining formulations have been suggested for use as topical disin-fectants and anti-bacterial agents that can be applied topically, orally orto animals such as cattle, birds or fish (Bischoff et al., 2015b).
In summary, liamocins are the only polyol lipids known to be
secreted by multiple strains of Aureobasidium in the ascomycetephylum, probably as a tool to adapt to leaf surface interactions.Liamocins are the most studied fungal polyol lipids and has promisinganticancer and antibacterial activities. At this point, research needs tofocus on determining the biosynthetic pathway and understanding thekey enzymes and how they are regulated in order to control its pro-duction. Knowledge of the metabolic pathway will also provide scien-tists with additional tools to create and tailor molecules that can serveas selective antimicrobials targeting particular applications in differentindustrial settings. Research also needs to be done to identify optimalproduction and downstream processing variables at scale, in order toestablish a techno-economical evaluation for commercialization.
3. Polyol esters of fatty acids
3.1. Producing strains
The vast majority of the PEFA secreting strains, mostly belong togenus Rhodotorula, were isolated from plant surfaces/plant surface in-teractions (Table 1) in agreement with previous studies (Ruinen, 1963).PEFA secretion by species of Rhodotorula was first reported by di Menna(1958). They isolated the red yeast strain 2 K53 from the leaves ofpasture grasses in the North Island of New Zealand, deposited it in theCBS yeast collection in the Netherlands as strain number CBS 2826, anddesignated it as the type strain of species Rhodotorula graminis. DiMenna reported that intra- and extracellular fat drops were present incultures, and the extracellular lipid was visualized as fat drops with asize similar to the cells, more prominent when grown on potato glucoseagar (di Menna, 1958). Two years later, Deinema quantified intra- andextracellular lipids produced by cultures of Rhodotorula graminis CBS3043, isolated from the leaf surface of citrus plants from Bogor, In-donesia (Deinema and Landheer, 1960), and Rhodotorula glutinis CBS3044 (Deinema, 1961), isolated from the leaf surface of Desmodiumrepens grown in tropical glasshouses at Wageningen, The Netherlands.These two yeasts produced up to 4 g/L intracellular lipid plus up to 2 g/L extracellular lipid.
In 1963, J. Ruinen studied 65 yeast strains isolated from Bogor(Indonesia), Paramaribo (Suriname) and Adiopodoumé (Ivory Coast),and reported the production of extracellular lipids from three morestrains of Rhodotorula glutinis, namely strains 14, 53 and 59, isolatedfrom Citrus aurantium, Theobroma cacao L. and Randia malleifera, re-spectively (Ruinen, 1963). The same publication reported secretion ofextracellular lipids of two strains of Rhodotorula mucilaginosa: strain 43and 49 (Ruinen, 1963). The extracellular composition of the lipids ofthe yeasts used in the study and the corresponding yields were notreported.
Two studies in 1964 (Ruinen and Deinema, 1964; Tulloch andSpencer, 1964) began to shed light into the structures of the intra- andextracellular lipids from R. glutinis. The first one published by Ruinenand Deinema (1964) revealed the fatty acid profile of both intracellularand extracellular lipids from R. glutinis strains CBS 3044 and CBS 4648,the latter isolated from cacao leaf surface in Ghana (Table 1). At thattime, they were able to identify the presence of six carbon polyols ascomponents of the extracellular lipids of both strains. The second study,published by Tulloch and Spencer (1964) reported a more detailedstructural analysis of the extracellular lipids produced by two Rhodo-torula glutinis strains, namely CBS 4648 and 16A8, and Rhodotorulagraminis strain 6CB. The authors reported that the extracellular lipidscontained 3-D-hydroxypalmitic and 3-D-hydroxystearic acids. R. glutinisstrain 16A8 and R. graminis strain 6CB were isolated from flowers ob-tained from several localities in Canada.
In the years since these studies were published, numerous taxo-nomic revisions have been made, including changes in yeast namesbetween older literature and recent publications (Kurtzman et al., 2011;Wang et al., 2015). Recent molecular discrimination of Rhodotorulaspecies using the Internal Transcribed Spacer (ITS) (White et al., 1990)
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and the nuclear large subunit (26S) ribosomal DNA partial sequences(D1D2) (Kurtzman and Robnett, 1998) revealed, for example, thatmany yeast strains previously identified as R. glutinis or R. graminis werereclassified as the close relative Rhodotorula babjevae. Strain CBS 4648used by Tulloch, which was originally identified as Rhodotorula glutinis,is now identified as Rhodosporidiobolus azoricus in the online CBS straincatalog (http://cbs.knaw.nl). In a similar fashion, R. glutinis strain IIP-30 reported by Johnson et al. (1992) to secrete surface active extra-cellular lipids is now identified in the ATCC catalog (https://www.atcc.org/) as Rhodotorula toruloides ATCC 204091 (see Table 1).
After the study of Johnson et al. (1992), research on PEFA entered astandby period, except for a few reviews (Amaral et al., 2010; Campos-Takaki et al., 2010; Van Hamme et al., 2003) citing previous work. Nofurther studies advancing the understanding of PEFA and their produ-cing strains were published until a recent detailed structural analysis(Cajka et al., 2016) as well as an in-depth screening for new producingstrains (Garay et al., 2017b). In the latter study, sixty-five yeast strainsin the order Sporidiobolales were screened, identifying nineteen strainscapable of secreting specific PEFA in yields ≥ 1 g/L (Table 1). Twentyadditional strains were found to secrete trace amounts of PEFA (Garayet al., 2017b). Rhodotorula paludigena UCDFST 81-492, R. aff. paludigenaUCDFST 81-84, Rhodotorula diobovata UCDFST 08-225 and Rhodotorulakratochvilovae UCDFST 05-632 are four new species reported to be alsocapable of producing and secreting> 1 g/L PEFA. Garay et al. (2017a,2017b) also identified that the phenotype within the same species canchange dramatically. For example, we found that R. babjevae strainUCDFST 04-877 secretes up to 8.6 g/L, while R. babjevae UCDFST 68-916.1 produces only 1.1 g/L when cultivated under identical condi-tions.
3.2. Structures of PEFA
The structures of nineteen different PEFA molecules have beenelucidated (Cajka et al., 2016, Garay et al., 2017a, 2017b), summarizedin Fig. 1. PEFA contain an acetylated (R)-3-hydroxyacyl tail group es-terified to a polyol head group through the carboxyl end of the fattyacid. The most common polyols present in PEFA are D-arabitol or D-mannitol (Cajka et al., 2016), and in one report low amounts of xylitolwere also observed (Tulloch and Spencer, 1964). All the PEFA-produ-cing strains currently known secrete mixtures of PEFA molecules,containing either D-mannitol as predominant head group, similaramounts of D-mannitol and D-arabitol head groups, or D-arabitol aspredominant head group. Unlike the non-acetylated polyol hydroxygroups in liamocins produced by Aureobasidium species, those samegroups in PEFA produced by Rhodotorula show different degrees ofacetylation (see Fig. 1). The acetylated (R)-3-hydroxyacyl tail group inPEFA is saturated, and the chain length is commonly 16 to 18 carbonslong, but small amounts of PEFA with chain lengths of 14 or 20 carbonshave also been reported (Cajka et al., 2016; Garay et al., 2017b).
The seven most abundant and recurrent PEFA across the differentPEFA-producing strains appear in Table 5. Ratios of these PEFA varydepending on the type of yeast strain. As an example Fig. 3 shows ex-tracted ion chromatograms of PEFA from two strains obtained by the
analysis of their extracts using reversed-phase liquid chromato-graphy–mass spectrometry operated in positive electrospray ionization.These PEFA contain either a fully acetylated polyol head group, or apolyol head group with one free hydroxy head group and the restacetylated. PEFA with polyol head groups containing two or more freehydroxy groups have been identified, but in less abundance and infewer yeasts. PEFA profiles can vary drastically depending on the yeastspecies and strain (Garay et al., 2017b), and to a lesser extent on thegrowth conditions (Deinema, 1961) and carbon sources used (Deinema,1961). Rhodotorula. aff. paludigena UCDFST 81-84 produces PEFAwith> 97% D-arabitol head groups (mainly C29H50O11, C31H52O12 andC31H54O11, see Table 5) (Garay et al., 2017a), whether grown on glu-cose (Garay et al., 2017a) or glycerol (Garay, 2017). In contrast, R.babjevae UCDFST 04-877 produces mixtures of PEFA containing D-ara-bitol and D-mannitol, and the profiles can change depending on the typeof carbon substrate used. When grown in varying amounts of glucose,the major PEFA molecules produced by R. babjevae UCDFST 04-877 areC31H52O12, C33H56O12 and C34H58O13 (Table 5). Conversely, whengrown in molasses, the major PEFA molecules produced are C32H54O13,C34H56O14 and C34H58O13, all of them with D-mannitol head groups.Different strains from the same species can also have marked PEFAprofile variation (Garay et al., 2017b). For example, the three mostabundant PEFA molecules in R. babjevae strain UCDFST 98-916.1 areC34H56O14, C31H54O11 and C33H56O12 (see Table 5) when grown inglucose, differing markedly from PEFA produced by R. babjevaeUCDFST 04-877 (Garay et al., 2017b).
Unlike the head groups of liamocins produced by Aureobasidium, the
Table 5Description of the seven most recurrent PEFA identified in Rhodotorula yeast. Using RPLC–ESI(+)-MS PEFA are detected as ammonium and sodium adducts. Peak numbering correspondsto peaks present in Fig. 3.
Peak no. Chemical formula m/z [M + NH4]+ m/z [M + Na]+ Polyol head group Acetylated (R)-3-hydroxyacyl tail Number of acetylations in polyol headgroup
1 C29H50O11 592.3697 597.3251 D-arabitol C16:0 32 C32H54O13 664.3908 669.3457 D-mannitol C16:0 43 C34H56O14 706.4014 711.3562 D-mannitol C16:0 54 C31H52O12 634.3803 639.3357 D-arabitol C16:0 45 C34H58O13 692.4221 697.3770 D-mannitol C16:0 46 C31H54O11 620.4010 625.3564 D-arabitol C18:0 37 C33H56O12 662.4116 667.367 D-arabitol C18:0 4
Fig. 3. RPLC–ESI-(+)-MS extracted ion chromatograms showing the most abundantPEFA isolated from (A) Rhodotorula diobovata UCDFST 08-225, and (B) Rhodotorula kra-tochvilovae UCDFST 05-632. Peak numbering corresponds to the different PEFA depictedin Table 5.
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head groups of PEFA produced by Rhodotorula species vary little whenproduced on different carbon sources. When R. glutinis and R. graminiswere grown using glycerol as the main carbon source, the resultingPEFA were virtually devoid of glycerol head groups (Deinema, 1961;Stodola et al., 1967). Studies conducted in our laboratory using R.babjevae UCDFST 04-877 and R. aff. paludigena UCDFST 81-84 obtainedsimilar results: all the PEFA detected had either D-arabitol or D-mannitolas head groups (unpublished data). These findings are in contrast withthe observations made by Price et al. (2017), in which growth of A.pullulans NRRL 50380 using glycerol as the carbon source resulted inliamocins with glycerol as the major head group (Price et al., 2017).
In 1964, Tulloch and co-workers studied the PEFA structures pro-duced by R. graminis strain 6CB, and R. glutinis strains CBS 4648 and16A8 (Tulloch and Spencer, 1964). They first used acid methanolysis torelease free polyols, methyl esters of 3-hydroxy acids and methylacetate (Tulloch and Spencer, 1964). The polyols were further analyzedas the acetates by GC–FID, while the 3-hydroxy acid was analyzed via acombination of techniques comprising reduction with red phosphorousand hydrogen iodide to discard the presence of branched carbon chains,identification of the carbon numbers on a silicone column and separa-tion of C16 and C18 methyl esters by countercurrent distribution(Tulloch and Spencer, 1964). The 3-hydroxy position was determinedvia release of myristic acid after chromic acid oxidation. However, thestudy did not reveal certain aspects of the chemical structures: thepoints of substitution of the acetyl groups, and the nature of the linkagebetween the polyol and the 3-hydroxy acid (Tulloch and Spencer,1964). Furthermore, the study did not shed light on the individualstructures and amounts in the mixture of similar PEFA molecules se-creted by these strains. The study of PEFA structure laid virtually dor-mant for several decades, until recent in-depth analysis by our group(Cajka et al., 2016).
After the discovery of new PEFA-secreting yeasts from the PhaffYeast Culture Collection (phaffcollection.ucdavis.edu), a new series ofstructural studies was performed that revealed the final aspects of PEFAstructure. In 2016, Cajka et al. (2016) used a multiplatform massspectrometry approach involving four different strategies. The firststrategy involved hydrolysis/neutralization of PEFA and analysis usinghydrophilic interaction chromatography–electrospray ionization in ne-gative mode-mass spectrometry (HILIC–ESI(−)-MS) in an untargetedmanner leading to annotation of polyols and hydroxy fatty acids inhydrolyzed extracts. The second strategy involved hydrolysis of PEFAand derivatization of the polyol and the 3-hydroxyacyl group followedby analysis via GC–electron ionization-MS (GC–EI-MS) with the ap-propriate standards to confirm the nature of their structures. The thirdstrategy encompassed analysis of the native PEFA molecules, by meansof a reversed-phase liquid chromatography–ESI in positive ion mode-tandem mass spectrometry (RPLC–ESI-(+)-MS/MS) to fragment themolecules and then reconstruct them using specialized software forstructural elucidation and the information obtained from the previoustwo strategies. The fourth strategy involved derivatization and chiralseparation GC–EI-MS to account for the chirality (stereochemical ana-lysis) of the different PEFA components (D- vs. L-polyols and (R)- vs. (S)-3-hydroxy fatty acids). This study revealed the remaining aspects of thechemical structure: the degrees of substitution of the acetyl groups inthe polyol head groups for each individual molecule, the nature of thelinkage between the polyol and the 3-hydroxy acid, the chain length ofthe 3-hydroxy acid for each PEFA molecule, and the individual struc-tures and relative abundances of PEFA molecules secreted by eachstrain (Cajka et al., 2016; Garay et al., 2017b).
3.3. Growth conditions, substrates and scale up productivities
PEFA production occurs when the yeast switches from the ex-ponential to the stationary phase, presumably triggered by the deple-tion of an essential nutrient in the growth media such as nitrogen. Thisphenomenon is characteristic of oleaginous yeast, and has been
thoroughly studied for intracellular lipid production, typically in theform of triacylglycerols (Garay et al., 2014; Sitepu et al., 2013) or sterolesters (Schweizer, 2004). The amount of carbon present in the growthmedium (typically in the form of a sugar such as glucose) is in highproportions compared to nitrogen such that the C:N ratio is preferablyabove 30. For a comparative view of each media, including the con-centrations of each ingredient, see Appendix A. For example, Deinema,1961 (Deinema, 1961) used a simple media containing 35–40 g/Lglucose, 1 g/L (NH4)2SO4, 1 g/L KH2PO4 and 0.2 g/L MgSO4·6H2O (C:Nratio = 66–75) for PEFA production from R. graminis, CBS 3043 and R.glutinis CBS 3044 (see Table 1), obtaining PEFA yields of 2.0 g/L and0.5–1.0 g/L respectively. She then examined the effect of nitrogen inPEFA production using 0.85, 1.55 and 2.2 g/L (NH4)2SO4 as sole ni-trogen source (equivalent to C:N ratios of 89, 49 and 34 respectively)and obtained 2.5, 0.5 and 0 g/L PEFA, respectively, suggesting thathigher C:N ratios were required for optimal PEFA production. She wasalso able to show that yeast extract as nitrogen source would give si-milar or higher PEFA yields compared to ammonium sulfate in the caseof R. graminis CBS 3043 and R. glutinis CBS 3044 respectively, keepingthe rest of the incubation conditions and media ingredients constant.We have varied the sugar and the sugar concentration (thus the C:Nratio) in our studies. The C:N ratios used for PEFA production in ourlaboratory varied between 50:1 and 300:1 (Cajka et al., 2016; Garayet al., 2017a; Garay et al., 2017b; unpublished data).
Deinema studied the effects of temperature in PEFA production forR. graminis CBS 3043 (Deinema, 1961) by cultivating the yeast at 18 °Cand 29 °C. She obtained almost twice the amount of PEFA at 18 °C(4.0–4.5 g/L) compared to 29 °C (2.0–2.2 g/L). Studies identified anoptimum pH range from 5.5–6.5 for PEFA production. Ammoniumsulfate caused a sharp pH fall in the culture media with time, requiringaddition of sterile KOH solution to keep pH values above 4.5, whereasyeast extract caused a milder pH fall between 4.5 and 5.5. Work done inour lab has shown that the ideal temperature for growth and PEFAproduction is between 24 °C and 27 °C, and using the inherent pH of themedium (medium A, pH = 6.5, described below) is optimal for PEFAproduction, since the natural pH decreases to 6.0–6.3 (Garay et al.,2017a).
The medium most commonly used for PEFA production is medium A(Garay et al., 2017b), developed by Suutari et al. (1993) to induce highlipid accumulation in oleaginous yeast (Holdsworth et al., 1988; Suutariet al., 1993). Medium A has a combination of NH4Cl as inorganic ni-trogen source and yeast extract as organic nitrogen source. The mediumis buffered at pH 6.5 with Na2HPO4 and KH2PO4, and contains thefollowing micronutrients: CaCl2, FeCl3·6H2O, ZnSO4·7H2O, MnSO4·H2O,CuSO4 and Co(NO3)2. Standard medium A contains 30 g/L glucose(Holdsworth et al., 1988); R. babjevae UCDFST 04–-877 and R. aff.paludigena UCDFST 81-84 produced PEFA when grown in D-glucose,sucrose, molasses, plant based hydrolysates and glycerol (Garay, 2017).Rhodotorula aff. paludigena UCDFST 81-84 produced 3.3 g/L PEFA from50 g/L molasses, 7.3 g/L PEFA from 50 g/L glycerol, and 8.4 g/L PEFAwhen grown in hydrolysate containing the equivalent of 80 g/L sugarsat shake flask scale (Garay, 2017).
These values are comparable to results of PEFA production byothers. When Rhodosporidiobolus azoricus (previously identified asRhodotorula glutinis) CBS 4648 was grown in basal media with eitherglucose, mannitol, arabitol or glycerol as main carbon source, yieldswere higher for mannitol and arabitol, followed by glucose and lastlyglycerol (Stodola et al., 1967). In scale-up studies using 7 L benchtopbioreactors (Garay, 2017) we obtained yields as high as 19 g/L from100 g/L glycerol. Up to 34 g/L PEFA was produced in a fed batchbioreactor, using an initial sucrose concentration of 100 g/L plus ad-dition of 50 g/L sucrose at 48 h in 4 L bioreactors after 7 days of in-cubation (Table 2). PEFA yields using sucrose were higher than thoseobtained with glucose (Table 2). Sucrose is an attractive raw materialbecause it is produced commercially as low-cost sugar cane syrup. Asuccessful economic model for biosurfactant production minimizes raw
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material cost, which in other EFGB like SL accounts for up to 89% of theoperating cost (Ashby et al., 2013). A mixture of hydrophobic (e.g. oleicacid or vegetable oil) and hydrophilic carbon sources (e.g. glucose) isrequired for SL production by the non-oleaginous yeast strains (e.g.Starmerella bombicola). Hydrophobic carbon sources are more ex-pensive, and require further downstream processing steps to removethem from the final product. PEFA producers do not require the use ofhydrophobic carbon substrates for their production because the yeaststhemselves are oleaginous, making PEFA producers attractive candi-dates for commercialization.
As mentioned previously for the case of liamocins, we and othershave observed that aeration is a key factor for PEFA production(Deinema, 1961; Garay et al., 2017a). PEFA production in shake flaskswas superior when using a culture to volume ratio of 1:5 or less, baffledflasks over smooth flasks, and foam stoppers rather than foil. Un-published comparative experiments involving cultivation of R. babjevaeUCDFST 04-877 in two different growth conditions: baffled shake flaskswith foam stoppers (high aeration) and smooth bottomed shake flaskssealed with foil (low aeration), translated into PEFA production of up to8 g/L in the former conditions, while no PEFA was recovered in thelatter conditions. This result suggested that aeration is important forPEFA production. Similar observations were obtained elsewhere(Deinema, 1961). Aeration has also been reported to be indispensiblefor growth of other EFGB like SL (Van Bogaert et al., 2007b) and MEL(Adamczak and Bednarski, 2000). Based on these observations, weperformed bioreactor studies with high dissolved oxygen levels of 40%saturation (Cajka et al., 2016; Garay et al., 2017a), achieved by bub-bling air at 0.5 void volumes per minute and looping agitation withdissolved oxygen values to keep them constant. The inoculum used forPEFA production usually involves plating the yeast from frozen stocks,then creating a one, two or three stepped seed train cell culture to thedesired scale, supplying each step with the proper aeration, as de-scribed (Garay et al., 2017a, 2017b).
PEFA recovery requires a solvent extraction step, using nonpolarsolvents. The most common solvents cited in literature are ethyl acetate(Cajka et al., 2016; Garay et al., 2017b) and ethyl ether (Deinema,1961). Higher yields of PEFA have been reported when extracting thefull culture rather than culture supernatant, just as observed for lia-mocins. Solvent is removed by evaporation with a stream of air or ni-trogen (Deinema, 1961), or using reduced pressure at room tempera-ture (Cajka et al., 2016; Garay et al., 2017b).
PEFA can be co-produced along with triacylglycerols in differentratios, depending on the yeast strain used (Garay et al., 2017a). Rho-dotorula aff. paludigena UCDFST 81–84 was able to secrete and accu-mulate 20.9 ± 0.2 g/L PEFA and 8.8 ± 1.0 g/L triacylglycerol,equivalent to an overall 24% conversion when grown in aerated fedbatch bioreactors using an initial glucose concentration of 100 g/L plusaddition of 50 g/L glucose at 48 h. The yeast strain R. babjevae UCDFST04–877 secreted 11.2 ± 1.6 g/L PEFA and 18.5 ± 1.7 g/L tria-cylglycerol, equivalent to an overall 22% conversion of glucose undersimilar conditions (Garay et al., 2017a). Secretion of a higher valuelipid is more desirable from an economic standpoint, making Rhodo-torula aff. paludigena UCDFST 81-84 a suitable candidate for furtheranalysis and commercialization.
3.4. Physiological role of PEFA
PEFA may play a role in yeast survival in extreme environments.Deinema (1961) conducted a series of studies using R. graminis CBS3043 and R. glutinis CBS 3044 in shake flask cultures left for up to tenmonths of incubation. The results show that cells remain viable as longas PEFA is present in the medium. PEFA was depleted at one to twomonths, and concomitantly viability decreased sharply for both strains.Deinema concluded that PEFA could be considered as a carbon reservefor yeast survival, and that the dead cells were not consumed by livingones (Deinema, 1961).
All basidiomycetous strains with known origin indicated in Table 1,except R. babjevae UCDFST 67-102 and UCDFST 08-225, were isolatedfrom either the phyllosphere or from other plant surface interactions'sources, suggesting that PEFA could play a role for survival in suchenvironments. Other glycolipid biosurfactants, like SL, are produced bystrains that were also isolated from similar environments. For example,Starmerella bombicola, a well-known SL producer, was isolated fromhoney of Bombus sp. (bumblebee) (Rosa and Lachance, 1998), andRhodotorula bogoriensis, the only basidiomycete known to produce SL(Zhang et al., 2011), was isolated from the leaf surface of Randia mal-leifera. Both are SL producers. The former is an ascomycete and knownto produce up to 400 g of SL per L in the presence of hydrophobicsubstrate added to the growth media (Pekin et al., 2005), while thelatter is a basidiomycete also known to produce a slightly different typeof SL.
Deinema (1961) showed that R. graminis CBS 3043 and R. glutinisCBS 3044 had extracellular lipolytic activity using three different ex-perimental approaches (for details on the methods used to determinelipolytic activity, see Deinema, 1961). Considering that: a) these yeastswere isolated from the phyllosphere, b) they secreted PEFA, and c) theyalso displayed extracellular lipolytic activity, Ruinen (1963) hypothe-sized that PEFA could play a role in the breakdown of cutin and even ofthe cuticle proper. She suggested that this modification in the yeastimmediate environment might enhance the permeability of the outerepidermis wall and ultimately lead to microbial growth (Ruinen, 1963).These findings led to the hypothesis that EFGB could be a feature morecommon than thus far considered in yeast whose habitat is the phyl-losphere (Ruinen, 1963). Further research is required to test these hy-potheses.
Certain species of Rhodotorula have also been described as potentialbiocontrol agents for plant pathogens. For example, R. glutinis (Yanet al., 2014) and R. paludigena (Wang et al., 2010) have been studied aspotential biocontrol agents against Alternaria alternata, a cherry tomatopathogen. A recent publication by Sen et al. (2017) reveals that gly-colipids secreted by a Rhodotorula babjevae strain have anti-fungal ac-tivity against a few important fungal plant pathogens. The glycolipidwas described as sophorolipid, which has very similar molecular for-mula but different chemical structure from PEFA; it is not clear whetherthis strain of R. babjevae produces sophorolipids rather than PEFA, or ifthe chemical analysis was not sufficiently detailed to distinguish thechemical structure.
3.5. Biosynthesis of PEFA
The full PEFA biosynthetic pathway is as yet unknown. However,since PEFA synthesis has been shown to occur in concomitance withtriacylglycerol synthesis (Garay et al., 2017a), and high levels of acetylCoA have been reported to occur right before secretion of PEFA(Deinema, 1961), it is probable that formation of the fatty acid portionof PEFA undergoes a similar biosynthetic pathway to that of tria-cylglycerol. In triacylglycerol biosynthesis, the primary building blockis acetyl-CoA. An important pool of acetyl-CoA is thus needed to movetriacylglycerol biosynthesis forward, and is likely the case for PEFAbiosynthesis as well. An intracellular pool of acetyl-CoA for triacylgly-cerol synthesis occurs in the cytosol by translocating citrate from themitochondria to the citosol via citrate/malate translocase (Ratledge andWynn, 2002), followed by cleavage in the cytosol by ATP citrate lyase(Garay et al., 2014). Acetyl-CoA is transformed into malonyl-CoA andthen fed to the fatty acid synthase I (FAS I) in order to synthesize thefatty acyl moiety present in triacylglycerol. The fatty acid moiety inPEFA could be synthesized in a similar fashion. The fate of the fatty acidchain leaving FAS I towards synthesis of PEFA is unknown. In otherEFGB such as SL, the fatty acid is hydroxylated by a CYP52 cytochromeP450 (Van Bogaert et al., 2009) coupled to a NADPH cytochrome P450reductase (Van Bogaert et al., 2007a), which are found to be bound tothe endoplasmic reticulum membrane. There are no reports on how the
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3-hydroxy fatty acids in the Rhodotorula strains are synthesized, andthus it is left to discover whether the mechanism is similar to the pro-posed for Aureobasidium strains (Section 2.5), except for one last in-complete reductive cycle, leaving a fatty acid of 16 or 18 carbons with a3-hydroxy group as a result. A recent genetic study revealed that no PKSgenes were present in several basidiomycete yeasts from the Puccinio-mycotina subphylum (Lackner et al., 2012), suggesting that the hy-droxylation of the fatty acid would likely be through a mechanismdifferent from PKS. The genomes of R. babjevae UCDFST 04-877 and R.aff. paludigena UCDFST 81-84 were sequenced and annotated, and noPKS genes were found, in agreement with previous observations (un-published data).
If PEFA is synthesized from glucose as carbon source, there must bea reductive step involved for the synthesis of mannitol or arabitol.Experiments performed in the 1960s revealed the existence of mannitoldehydrogenase activity in cell free extracts from different Rhodotorulaglutinis strains. The results pointed to two different types of enzymes,one dependent on NAD+ and the other dependent on NADP. Thesecond turned out to be specific to the PEFA-secreting strains. R. glutinisPEFA secreting strains had also positive growth in mannitol and ara-bitol, with higher yields of PEFA (Stodola et al., 1967). This evidencesuggests that glucose can be reduced to mannitol for PEFA synthesis.
3.6. Properties, uses and applications of PEFA
Previous studies have suggested that PEFA properties have potentialfor biotechnology application. The appearance of PEFA is similar tohoney: viscous, with color varying from golden/orange to dark, reddishbrown. PEFA is denser than water, with a density around 1.08 g/mL(Garay et al., 2017b). PEFA has been visualized under the microscopeas early as 8 (Deinema, 1961) or 17 h (Garay et al., 2017a) after theinception of fermentation, presenting as amorphous droplets thatfluoresce profusely when stained with Nile red and soon exceed the sizeof the cells as fermentation progresses.
The physicochemical properties of PEFA suggest their function asextracellular surface active agents with the potential to act as techno-logically valuable biosurfactants. For example, PEFA mixtures producedby R. aff. paludigena UCDFST 81-84 showed antifoam activity compar-able to commercial antifoaming agents used in the brewing industry(data not shown). When mixed with water, the surface tension values ofPEFA produced by R. aff. paludigena UCDFST 81-84 and R. babjevaeUCDFST 04-877 in our laboratory decreased with PEFA concentration,attaining minimum values of 33.3 and 30.4 mN/m respectively(Table 3). In comparison, surface tensions of SL have been reported torange between 33 and 38 mN/m (Chandran and Das, 2011; Chen et al.,2006; Solaiman et al., 2004). Solubility limits in water ranged from1.3–1.5 mg/L for R. aff. paludigena, and was 0.9 for R. babjevae UCDFST04-877. PEFA from these strains were highly soluble in organic solvents(e.g. ethyl acetate), and promote the formation of water-in-oil emul-sions in water/octane mixtures (Xu, 2017).
To date, it is unknown whether PEFA has antibacterial or anticanceractivities. The structural similarities between liamocins and PEFAsuggest that they could have such activities, but research is needed toprove this hypothesis.
4. Future steps on extracellular fungal polyol lipid research
A competitive advantage for commercialization of both liamocinsand PEFA is that both Aureobasidium and Rhodotorula species synthesizea variety of high value molecules simultaneously with liamocins andPEFA respectively. Such complex growth properties argue for their usein a biorefinery model in which multiple co-products can be valorizedfrom a single culture (Kamm and Kamm, 2007). For example, the acidicpolyester PMA, the exopolysaccharide pullulan, and liamocins are si-multaneously produced by Aureobasidium sp. cultivated in a relativelysimple bioreactor process with low cost carbon source such as
sugarcane syrup, which is rich in sucrose. The co-products can also berecovered relatively easily due to their extracellular nature and theirdiffering physicochemical properties. Manitchotpisit et al. (2012) pro-vided a lab scale proof of principle method by separating the threeextracellular co-products using careful centrifugation to separate thespent media (containing pullulan and PMA) from the cell pellet (con-taining liamocins). Pullulan was then precipitated from the spent mediaby first adjusting the spent media to pH 5 with CaCO3 and then adding95% ethanol. After removal of pullulan fraction by centrifugation, thePMA was separated by precipitation with 95% ethanol followed byincubation at 4 °C overnight and separation via centrifugation. Liamo-cins can then be recovered from the cell pellet through solvent ex-traction, as described in Section 2.3 (Manitchotpisit et al., 2011). Theyapplied this method to screen several A. pullulans strains, and thehighest producer of the three co-products was A. pullulans NRRL Y-12974 with 6.7, 19 and 1.9 g/L of PMA, pullulan and liamocins re-spectively. These values add up to a net production of 27.6 g/L productfrom 50 g/L glucose, representing a conversion of 0.552 g of totalproducts per gram of glucose. Different strains of A. pullulans produceddifferent weight ratios of PMA, pullulan and liamocins (Manitchotpisitet al., 2012). For example, strain CU 47 (NRRL 58558) produces aPMA:pullulan:liamocin ratio of 1.97:1.18:1, whereas strain RSU 7 cre-ates PMA:pullulan:liamocin in a 17.8:19.6:1 ratio. A. pullulans strainNRRL Y-12974 has a ratio of 3.52:10:1 since it is a high pullulan pro-ducer. The work done by Manitchotpisit et al., 2012 was a successfulproof of principle that the three co-products can be successfully sepa-rated and purified from a single A. pullulans culture. However, moreresearch is needed in order to assess the techno-economic feasibility forthis approach, focusing on the spectrum of productivities that help toadapt to commercial scenarios where demand and price of the differentco-products vary. During high demand of pullulan, production fromstrain NRRL Y-12974 would be more desirable, whereas during highdemand of liamocin, strain CU47 could be preferred.
All the PEFA-producing strains present in Table 1 also simulta-neously accumulate intracellular lipid in the form of triacylglycerol(Garay et al., 2016; Sitepu et al., 2013), and many of them also producecarotenoids (Frengova and Beshkova, 2009). The weight ratio of in-tracellular triacylglycerol to PEFA seems to be above one for all PEFAproducing strains present in Table 1 except for Rhodotorula aff. palu-digena UCDFST 81-84, which produces more PEFA per unit of in-tracellular lipid (Garay et al., 2017a). In a trial designed to demonstratemechanical extraction of the different lipid fractions, several liters of R.babjevae were cultivated in shake flasks, and PEFA was collected bycentrifugation, while the intracellular oil was extracted using a screwpress. The oil contained triacylglycerols with oleic acid as the majorfatty acid (Garay et al., 2016), making the oil a suitable precursor forbiodiesel. Carotenoid pigments were also present in the oil. The latterprocess would recover three high value lipid-based fractions with apurely physical downstream recovery process, involving no solvents,strong acids or bases.
The current yields obtained for both liamocins and PEFA are belowthose obtained for other EFGB produced at industrial scale. For ex-ample, SL produced by S. bombicola have been produced in yields of upto 400 g/L (see Table 2) (Daniel et al., 1998), roughly ten times morethan liamocins and PEFA. However, achieving these yields took decadesof intense research on growth optimization, scale up, strain manipula-tion, elucidation of the biosynthetic pathway and other types of re-search that allowed to transition from initial yields around 50 g/L(Gorin et al., 1961) to yields of> 400 g/L (Pekin et al., 2005). A similarapproach could be taken in the case of polyol lipids to achieve similaryields, incorporating the advantage of valorization of co-products.
Conflict of interest
The authors declare that they have no conflicts of interest.
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Acknowledgments
This work, including the efforts of L.A. Garay was funded by theNational Mexican Council of Science and Technology (CONACYT)[Fellowship number 291795]. This work, including the efforts of T.Cajka was funded by the National Institutes of Health [grant numberNIH HL113452 and grant number NIH DK097154], as well as the NIHinstrument funding [grant number NIH S10-RR031630]. This work,including the efforts of I.R. Sitepu, J. Xu, S.R. Dungan and K.L. Boundy-Mills was funded by award number 1 from the University of CaliforniaDavis Science Translation and Innovation Research (STAIR) program.All authors have agreed to submit this manuscript to BiotechnologyAdvances.
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.org/10.1016/j.biotechadv.2018.01.003.
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