how could haloalkaliphilic microorganisms contribute to biotechnology?
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How could haloalkaliphilic microorganisms contribute to biotechnology? 1
Baisuo Zhao1*
, Yanchun Yan1, Shulin Chen
2 2
1Graduate School, Chinese Academy of Agricultural Sciences, Beijing 100081, China 3
2Biological Systems Engineering, Washington State University, Pullman, WA 99164, USA 4
*Corresponding author: [email protected] 5
Tel: 86-10-82106784 6
Fax: 86-10-82106784 7
Running title: Haloalkaliphilic microorganisms contribute to biotechnology8
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9
Abstract: 10
Haloalkaliphiles are microorganisms requiring Na+ concentrations of at least 0.5 M and alkaline pH 9 11
for optimal growth. Their unique features enable them make significant contributions to a wide array of 12
biotechnological applications. Organic compatible solutes produced by haloalkaliphiles such as ectoine and 13
glycine betaine are correlated to osmoadaptation and may serve as stabilizers of intracellular proteins, salt 14
antagonists, osmoprotectants, and dermatological moisturizers. Haloalkaliphiles are an important source of 15
secondary metabolites like rhodopsin, polyhydroxyalkanoates, and exopolysaccharides that play essential 16
roles in biogeocycling organic compounds. These microorganisms also can secrete unique exoenzymes 17
including proteases, amylases, and cellulases that are highly active and stable in extreme halo-alkaline 18
conditions and can be used for the production of laundry detergent. Furthermore, the unique metabolic 19
pathways of haloalkaliphiles can be applied in the biodegradation/biotransformation of a broad range of toxic 20
industrial pollutants and heavy metals, wastewater treatment, and biofuel industry. 21
Key words: 22
Haloalkaliphile, Compatible solutes, Secondary metabolites, Exoenzymes, 23
Biodegradation/biotransformation, Biofuel industry24
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25
1. Introduction 26
Haloalkaliphiles are extremophilic microorganisms capable of robust growth in rigorous ecosystems 27
characterized by both high salinity and extreme alkalinity (Sorokin and Kuenen 2005a, 2005b). They are 28
found in all three domains of life: Archaea, Bacteria, and Eukarya. They are mostly confined to exceptionally 29
stable, naturally occurring hypersaline alkaline environments like soda lakes and soda deserts distributed in 30
various dry steppes and semidesert areas around the world. These include soda lakes of Siberian and Kulunda 31
Steppe in Russia, Mono Lake and Soap Lake in North America, Natron Lake and Magadi Lake in the East 32
African Rift Valley, Magadi Lake in Kenya, and the Wadi Natrun lakes in Egypt (Foti et al. 2007, 2008, 33
Horikoshi 1999, Sorokin and Kuenen 2005). Such extremely halo-alkaline habitats are caused by the 34
combined effects of topography, geochemistry, and climate (Grant and Mwatha 1989, Kulp et al. 2007, 35
Sorokin et al. 2008). Topography is responsible for hydrological phenomena leading to a permanently 36
enclosed body of water. Geochemistry determines the major ionic composition and total salt content of these 37
endorheic drainage basins. Climate may lead to the accumulation of salts when evaporation rates exceed 38
in-flow rates. Saline soda lakes are characterized by the presence of high levels of salts, where the cation is 39
Na+ and K
+ as well as anion is mainly Cl¯, CO3
2¯, HCO3¯, and SO4
2¯. A correspondingly low concentration of 40
both Mg2+
and Ca2+
cations results from carbonate precipitation. 41
Athalassohaline lakes containing high levels of sodium carbonate / sodium bicarbonate represent a 42
combined conditions of both a salinity exceeding that of sea water (~35 g/L) and up to saturation and a pH 43
value generally around 9 and potentially up to 11 (Shapovalova et al. 2008, Sorokin et al. 2011). Microbial 44
populations in such athalassohaline harsh habitats thrive to the extent that they can be observed without the 45
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aid of a microscope as different shades of red, purple, and green, representing archaea, photosynthetic purple 46
bacteria, and cyanobacteria, respectively. Hyper-saline and alkaline environments can alternatively result 47
from human industrial processes, such as those involving mineral ore, petroleum refining, pulp and paper, 48
textile preparation, leather tanneries, food and potato processing units, calcium carbonate kilns, and detergent 49
manufacture which generate effluents containing NaOH, Ca(OH)2, etc. (Alnaizy 2008, De Graaff et al. 2011, 50
Jones et al. 1998). 51
In general, microorganisms growing optimally at a NaCl concentration at or above 0.2 M, which differ 52
from natronophiles inhabiting in sodium carbonate brines, can be referred to as halophiles (Oren 2006, 53
Ventosa et al., 1998). Depending on the previous authors’ definitions, microorganisms growing optimally 54
either at or above 9, 9.5, or 10 are termed alkaliphiles by most of researchers although this definition is 55
disputable (Horikoshi 2006; Krulwich 2006). So far, an absolute definition for what constitutes 56
haloalkaliphiles has been elusive in the literature. For this review, the authors wish to focus on 57
microorganisms requiring for optimal growth at least 0.5 M total Na+ concentrations in the form of NaCl or 58
pH-mediating sodium carbonate / bicarbonate, and alkaline pH values of at least 9 (Table 1). 59
The term ‘haloalkaliphilic’ was earlier used to describe a novel extremophilic archaeon, Natronomonas 60
pharaonis DSM 2160T (formerly known as Natronobacterium pharaonis) originally isolated from biotopes of 61
Abu Gabara Soda Lake of Wadi Natrun in Egypt (Soliman and Trüper 1982), which possesses both a salinity 62
of 36 % and an alkaline pH of 11 (Imhoff et al. 1978). Subsequently, two haloalkaliphilic archaeal isolates, N. 63
pharaonis SP1T (DSM 3395
T) and Natronococcus occultus SP4
T (DSM 3396
T), were revived from the soda 64
lake Magadi in Kenya (Kamekura et al. 1997, Tindall et al. 1984). Since then, microorganisms that habitat in 65
the combined conditions of both high salinity and extreme alkalinity have been a topic of fascination to 66
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microbiologists. 67
The potential for the discovery and utilization of novel haloalkaliphiles and their products is immense, 68
as they provide unique physiological functions under extreme conditions that limit survivability of their 69
mesophilic counterparts. During the three last decades, there has also been an increase in interest and 70
investment to explore haloalkaliphiles as a precious source of organic compounds, stable exoenzymes, and 71
unique metabolic pathways with potential applications in various industrial processes. There are still many 72
interesting and unresolved questions with respect to how they may affect and even benefit human life. 73
Specifically, how could these haloalkaliphilic microorganisms contribute to biotechnology? The objective of 74
this review is therefore to summarize and focus on the recent developments pertaining to the potential for 75
haloalkaliphiles to ultimately contribute to the efforts of the biotechnology`industry in meeting expanding 76
market demand for haloalkaliphile-based applications. 77
2. Organic compatible solutes 78
The properties of haloalkaliphile-derived compatible solutes can be biotechnologically exploited for a 79
wide range of applications in cosmetology, medicine, and agriculture, etc. Ectoine and hydroxyectoine can 80
stabilize enzymes to prevent denaturation via desiccation and chemical agents, and to enhance protein folding 81
via chemical chaperones. They also can be used to enhance PCR performance (Schnoor et al. 2004) and DNA 82
microarray sensitivity and specificity (Mascellani et al. 2007). A low concentration of hydroxyectoine (10 to 83
25 mM) has been demonstrated to reduce DNA microarray background and improve hybridization efficiency 84
(Mascellani et al. 2007). Glycine betaine effectively protects against mutagenic compounds and 85
radiation-induced damage. It can protect plants against abiotic stress from salt, drought, heat, and chilling. 86
For example, glycine betaine can be exogenously applied to leaves and roots of agronomically imporant 87
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crops, like rice and potato, to increase tolerance to various stresses (Sakamoto and Murata 2000). 88
Compared to chemical synthesis, the haloalkaliphilic biosynthesis and subsequent extraction and 89
purification of compatible solutes like ectoine and glycine betaine are relatively easier and higher yielding. 90
The intracellular concentration of ectoine reached 235 µg/mg dry cells of Methylophaga alcalica M39T 91
grown at 1.6 M Na+ and pH 9.5 (Doronina et al. 2003b). Recently, a moderate haloalkaliphile, Methylophaga 92
lonarensis MPLT, intracellularly accumulated ectoine (149–197 µg/mg dry cells) as primary osmoprotectant 93
when grown under 1–1.5 M NaCl and pH 9–10 (Antony et al. 2012). Ectoine production by batch cultures of 94
the haloalkaliphilic Thioalkalimicrobium aerophilum AL 3T was proportional to extracellular Na
+ 95
concentration in mineral media (Banciu et al. 2005). Here, ectoine accounted for 8.7 % of total dry weight at 96
1.2 M Na+ and pH 10. Furthermore, the haloalkaliphilic Methylophaga murata Kr3
T synthesized ectoine, 97
where intracellular ectoine concentration increased from 40 µg/mg dry cells at 0.57 M Na+ to 160 µg/mg dry 98
cells at 1.60 M Na+ (Doronina et al. 2005). Interestingly, the intracellular ectoine concentration at a low 99
temperature of 4 ºC was two-fold than at a mesothermal temperature of 29 ºC, which is beneficial for 100
industrial processes that recover more products at short low temperature stress. When grown at high salt 101
salinity, the Kr3T cells could withstand heating, repeated freeze-thaw cycles, and lyophilization without 102
adding any cryoprotectants, indicating that ectoine could protect whole cells intact and its protein and DNA 103
against denaturation. The haloalkaliphilic Methylophaga natronica Bur2T also produced ectoine under 104
saline-alkaline conditions (Doronina et al. 2003a). In a 1.5 L continuous laboratory fermenter, 105
Thioalkalivibrio versutus ALJ 15 accumulated 7.5 %, and 9 % of total dry weight of glycine betaine as the 106
major compatible solutes at 2, and 4 M Na+, respectively (Banciu et al. 2005). The haloalkaliphilic 107
Desulfonatronospira thiodismutans ASO3-1 accumulated 16 % of total dry cell mass of glycine betaine at 3 108
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M total Na+ concentration (Sorokin et al. 2008a). Zhao and Wiegel investigated the accumulated compatible 109
solutes from the anaerobic halophilic alkalithermophilic bacterium Natranaerobius thermophilus 110
JW/NM-WN-LFT that grew optimally at 3.5 M Na
+, pH
55ºC 9.5, and 53 ºC. When grown at 52 °C and pH
55ºC 111
9.5, and when extra-cellular Na+ concentration was increased from 3.3 M to 4.5 M, the intracellular 112
concentration of glycine betaine in N. thermophilus increased more than three-fold from 0.437 M to 1.321 M. 113
However, the haloalkaliphiles are less efficient producers since they have to spend additional energy for pH 114
homeostasis when comparing with the neutraphilic halophiles for compatible synthesis production. Therefore, 115
publications involving hydroectoine, glutamate and other compatible solutes from haloalkaliphiles are limited 116
and to date have not yet practically investigated their biotechnological potential. 117
3. Secondary metabolites 118
Haloalkaliphilies can produce secondary metabolites of tremendous importance to biotechnology. These 119
include rhodopsin and the biopolymer of polyhydroxyalkanoate (PHA) and exopolysaccharides (EPS). 120
Haloalkaliphile-derived retinal proteins like halorhodopsins (hR) have many potential applications for the 121
manufacture of photochromic molecular materials used for data storage, light-induced color changing, 122
holographic films, instant photoelectrical response, nonlinear optical capability, etc. Bivin and Stoeckenius 123
(1986) reported that thirty eight haloalkaliphilic bacteria isolated from alkaline salt lakes in Kenya and the 124
Wadi Natrun in Egypt mainly possessed two photoactive retinal pigments. One pigment had spectroscopic 125
properties similar to those of hR. The other pigment had kinetics very similar to those of a sensory rhodopsin 126
(sR). Two types of rhodopsins, hR and phoborhodopsin (ppR; also known as sensory rhodopsin II), were 127
found in the haloalkaliphilic N. pharaonis SP1T (Duschl et al. 1990, Hirayama et al. 1992). The hR is a 128
light-driven chloride pump that maintains the effective level of chloride anions in the cell and has an 129
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absorption maximum between 575 and 580 nm. The ppR has an absorption maximum occurring at 498 nm 130
and can be stably solubilized in several kinds of detergents (Hirayama et al. 1992).The mutant strain KM-1 131
generated from N. pharaonis DSM 2160T over produced ten-fold greater amounts of hR than that the 132
wild-type due to a point mutation from As-Asp324 to Asn in the bacteriorhodopsin activator homologues 133
(Ihara et al. 2008). 134
Polyhydroxyalkanoates (PHAs) that are accumulated intracellularly as energy storage compounds 135
comprise a large number of polyesters synthesized from renewable carbon sources like agriculture waste via 136
bacterial fermentation. Secreted haloalkaliphilic polymers can be considered as a raw material for 137
biodegradable plastics. Owing to their biocompatible and biodegradable characteristics, PHAs are materials 138
of great interest for pharmaceutical and clinical purposes, and are regarded as an alternative to 139
non-biodegradable plastics produced from fossil-based resources (Braunegg et al. 1998, Strazzullo et al. 140
2008). A PHA-producing haloalkaliphilic bacterium Halomonas profundus AT 1214T grew optimally at 3 % 141
(w/v) sea salts and at pH 8–9 (Simon-Colin et al. 2008). The PHA yield of strain AT 1214 T
was nearly 270 142
mg/L in sea salts broth. The chemical composition of the PHAs consisted of different proportions of 143
poly-3-hydroxybutyrate-co-3-hydroxyvalerate (PHV) and poly-3-hydroxybutyrate (PHB) depending on the 144
various carbon substrates used. Strain AT 1214 T
is thus a viable candidate for the cost-effective manufacture 145
of PHAs from by-products of many industries. A haloalkaliphilic bacterium Halomonas campaniensis 5AGT 146
also produced PHAs (Gambacorta et al. 2005). At 1 M NaCl and pH 9 in the presence of glucose as carbon 147
source, PHB production represented more than 10 % of the wet cell weight (Strazzullo et al. 2008). Several 148
haloalkaliphilic microbes belonging to the species of Halomonas campisalis could also accumulate PHAs 149
granules Strain MCM B-1027 accumulated 54 % of dry cell weight as PHB co-PHV when grown in a 150
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production-medium containing 1 % (w/v) maltose at 0.51 M NaCl and pH 9 (Kulkarni et al. 2010). The PHA 151
accumulated by strain MCM B-1027 is a biodegradable co-polymer of both PHB (96 mol %) and PHV (4 152
mol %), has a melting temperature of 166.5°C and tensile strength of 18.8 MPa, which renders its potential 153
application as a packaging material (Kanekar et al. 2011). 154
Exopolysaccharides (EPSs) from haloalkaliphiles have unusual characteristics and functional activities 155
that can be exploited for applications in the food, pharmaceutical, and cosmetics industries. Knowledge of the 156
factors influencing haloalkaliphilic EPS production is unfortunately still very limited. The EPS produced by 157
haloalkaliphilic M. murata Kr3T was composed of carbohydrate and protein moieties that could stabilize the 158
cells and prevent them from drying (Doronina et al. 2005). Strain Kr3T represents a promising candidate for 159
cost-effective large-scale production of EPS as the result of the ability to use methanol, methylamine, 160
trimethylamine, and fructose as carbon and energy sources under a wide range of pH and salinity. 161
4. Exoenzymes 162
Momentum has recently gained in the study of the behavior of active haloalkaliphile-secreted 163
exoenzymes in the coupled stringent factors of high salt and alkaline pH conditions. Exoezymes derived from 164
haloalkaliphile are stable, functional, and efficient biocatalysts in organic solvents because the extracellular 165
surrounding medium environment that they are secreted into resembles non-aqueous systems due to the 166
reduction of water activity by high salt concentration. Therefore, haloalkaliphile-secreted exoezymes are 167
further being considered for manufacturing and bioremediation processes because they have catalytic ability 168
not only at high salinity and alkalinity, but also at the low-water activity of non-aqueous solvent or surfactant 169
environments. The sustainable production of fuels, chemicals, biopolymers, and materials from renewable 170
biomass resources like agricultural waste may also require haloalkaliphilic exoezymes that can withstand the 171
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high salt and pH conditions of pretreatment and hydrolytic degradation steps. These may include proteases, 172
α-amylases, cellulases, and xylanases, etc (Table 2). With all things considered, compared to enzymes from 173
halophiles or alkaliphiles, literature pertaining to those from haloalkaliphiles so far is limited and the 174
following includes a review of such works. 175
4.1. Proteases 176
Proteases are enzymes that conduct proteolysis, beginning protein catabolism by hydrolyzing the peptide 177
bonds that link amino acids together in the polypeptide chain forming proteins. They are important 178
constituents of detergents and typically need to be active and stable in conditions of high salt and surfactant 179
concentrations, alkaline pH 9–11 that prevail under harsh washing conditions. Therefore, the 180
haloalkaliphile-derived proteases have recently received much attention for their superior stability under 181
otherwise destabilizing industrial conditions. 182
The potentially useful proteases from the haloalkaliphiles Bacillus species have been purified and 183
extensively characterized. Protease production and secretion from the haloalkaliphilic Bacillus sp. strain Vel 184
reached a maximum level of 410 U/ml at optimal growth conditions of 10 % (w/v) NaCl and pH 9 (Patel et al. 185
2006). The protease secretion occurred with an optimum at pH 8–9 and was 186 U/ml, 172 U/ml, and 158 186
U/ml at 1.71, 2.56, and 3.42 M NaCl, respectively. It was active at 0–0.17 M salt concentration, pH 8.5–12, 187
and was optimal at pH 10–11 (Gupta et al. 2005). The protease was stable in various surfactants (i.e. 0.1 % 188
SDS, 0.1 % Triton X-100, and 0.1 % Tween 80) and heavy metal ions (i.e. 5 mM Mn2+
, 5 mM Zn2+
5 mM 189
Mg2+
, and 5 mM Cu2+
). The stability of this protease at the poly-extremities of high pH, high salt and the 190
presence of detergent components and surfactants as well as metal ions makes this enzyme particularly 191
suitable for its application in detergent industries.192
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193
The crude serine protease was secreted at 0.185 mg/ml by the haloalkaliphilic Bacillus sp.AH-6 and was 194
exceptionally resistant, retaining 65 %, 50 %, and 95 % of its original activity after 24 h incubation in 8 M 195
urea, 4 M guanidine-HCl, and 2 M guanidine-HCl, respectively (Dodia et al. 2008a). This protease exhibited 196
catalysis and stability at pH 8–13 and 0–4 M NaCl with an optimum coinciding with pH 9–11 and 0.15–0.2 197
M NaCl, and this was slightly enhanced by 0.2 % Tween-80 and 0.05 % Triton X-100 (Dodia et al. 2008b). 198
The NaCl salt enhanced the enzyme’s substrate affinity, as reflected by the increase in the catalytic constant 199
(Kcat). Salt requirement for optimal catalysis also increased with temperature. Interestingly, this protease 200
could regain stability and functionality in the presence of NaCl after chemically denaturing (Dodia et al. 201
2008a). Such rare phenomenon can be further investigated to enrich the body of knowledge regarding protein 202
stability and denaturation. Furthermore, this serine protease was highly stable in oxidizing-reducing agents 203
and commercial detergents. These attributes suggest that this enzyme can be used in detergent applications to 204
remove proteinaceous stains and deliver unique benefits otherwise unobtainable with conventional detergent 205
technologies (Dodia et al. 2008). 206
An extracellular protease from the haloalkaliphile Salinivibrio costicola 18AGT was also purified and 207
characterized (Lama et al. 2005, Romano et al. 2005). It showed optimal activity at 60 °C in the presence of 208
both 0.34 M NaCl and pH 8, with 80 % of residual activity at pH 9. The protease was slightly activated by 209
denaturing agents such as SDS (0.1 %) and urea (6.0 M). Karan et al. (2011) reported that 38 haloalkaliphilic 210
bacteria isolated from Sambhar Salt Lake (India) were able to secrete haloalkaliphilic proteases. Four of these 211
strains named EMB1, EMB2, EMB3, and EMB4 exhibited 28, 37, 21 and 11 U/ml protease activities in 212
gelatin broth, respectively. The crude protease from strain EMB2 belonging to the Geomicrobium genus 213
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exhibited appreciable stability at NaCl concentrations up to 3.42 M (optimum at 0.85 M NaCl), temperature 214
up to 70 °C (optimum at 50 °C), pH 6–12 (optimum at pH 10), and up to 75 % (v/v) concentrations of various 215
organic solvents. For instance, the protease still retained 39 %, 91 %, 98 %, and 101 % of its original activity 216
in cyclohexane, ethanol, benzene, and n-dodecane after 72 h incubation, respectively. The protease also was 217
compatible with up to 2 % (w/v or v/v) of surfactants (SDS, CTAB, Tween 20, Tween 80, Triton X-100, and 218
Triton X-114) and up to 1 % (w/v) of commercial detergents, indicating its potential use for industrial 219
applications. 220
As proteases can not only catalyze hydrolysis but also synthesize peptide bonds, they may represent an 221
alternative to chemical synthesis. A chymotrypsin-like and solvent-tolerant serine protease (Nep) from the 222
haloalkaliphilic archaeon Natrialba magadii was purified and characterized (Giménez et al. 2000). The Nep 223
was active and stable at pH 8 in mixtures of 1.5 M Na+ and up to 30 % (v/v) aqueous-organic solvents that 224
included glycerol, dimethylsulfoxide (DMSO), N,N-dimethyl formamide, propyleneglycol, and dioxane 225
(Ruiz and De Castro 2007). Under the optimal combined conditions of 30 % DMSO, 1.5 M NaCl, and pH 8, 226
Nep catalyzed the synthesis of the tripeptide Ac-Phe-Gly-Phe-NH2 from Ac-Phe-OEt ester and Gly-Phe-NH2 227
amide substrates (Ruiz et al. 2010). The Nep gene from N. magadii was cloned and expressed heterologously 228
in Haloferax volcanii, which represented the first recombinant system for the secretion of a haloarchaeal 229
protease and resulted in high level production and activity of Nep protein that biocatalyzed peptide synthesis 230
under low-water activity conditions (De Castro et al. 2008). The high activity and stability of this 231
heterologously expressed protease make it a promising candidate for future applications (Ruiz et al. 2010). In 232
addition, a chymotrypsinogen β-like protease from N. pharaonis exhibited optimal activity at between 0.5 233
and 4 M NaCl, pH 10, and 61°C (Stan-Lotter et al. 1999). Another serine protease that had chymotrypsin-like 234
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activity from N. occultus during the stationary phase was purified, and its activity and stability depended on 235
high salt concentrations of 1–2 M NaCl, a broad pH range of 5.5–12, and rather thermophilic conditions with 236
an optimal activity at 60 °C in the presence of 1–2 M NaCl (Studdert et al. 1997, Studdert et al. 2001). 237
4.2. α-amylases 238
α-amylases are enzymes that randomly hydrolyze starch molecules, and their demand is currently 239
increasing because of its great potential to be extensively applied in various biotechnological industrial 240
processes involving starch liquefaction for biofuels and the manufacture of detergent and pharmaceuticals. 241
However, the α-amylases typically isolated from non-haloalkaliphiles do not meet industrial requirements due 242
to their inability to withstand harsh inhibitory industrial conditions of high salt, surfactant, and detergent 243
concentrations. Therefore, isolation and exploration of novel haloalkaliphile-derived α-amylases with 244
desirable halo-, alkaline-, and thermo-stability is crucial to meet industrial demands. 245
The first reported haloalkaliphilic extracellular α-amylase was derived from Natronococcus sp. strain 246
Ah-36, an archaeon from Kenyan soda lake of Lake Magadii which could tolerate a wide salinity (1.4 M 247
NaCl–saturation), pH (8–10), and temperature (20–55°C) (Kobayashi et al. 1992). When strain Ah-36 was 248
aerobically cultivated at 3.5 M Na+ and pH 9, α-amylase activity of 0.01 U/ml was detected at 70 h in the 249
culture broth and reached a maximum of 0.12 U/ml after 90 h. This enzyme was active at 1–5 M NaCl and 250
was most stable at pH 6–8.6 with a maximum at pH 8.7 and 2.5 M NaCl. It cleaves α-1, 4 linkages 251
endolytically and hydrolyzes soluble starch, amylose, amylopectin, glycogen, and γ-cyclodextrin to produce 252
maltotriose of α-configuration as the major product. Therefore, biotechnological applications of α-amylase 253
include the treatment of saline water or waste solutions containing starch residues at halo-alkaline and 254
poly-stress conditions. Prakash et al. (2009) reported two kinds of the extracellular α-amylase, designated as 255
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α-amylase I and α-amylase II, optimally produced by the strain Chromohalobacter sp. TVSP 10 when grown 256
on 0.5 % rice flour at 3.4 M Na+ and pH 9. These were halo-alkalistable and moderately thermophilic 257
α-amylases with maximal activity at 0–3.41 M NaCl, pH 9, and 65°C that efficiently hydrolyzed 258
carbohydrates to yield maltotetraose, maltotriose, maltose, and glucose as end products. Recently, a 259
haloalkaliphilic α-amylase that was stable in surfactants, oxidants, and detergents was extracellularly secreted 260
from the actinomycete Saccharopolyspora sp. A9 at 1.9 M salinity and pH 11 (Chakraborty et al. 2011). This 261
enzyme hydrolyzed starch primarily to glucose, maltose, and maltotriose. It retained 72 % and 33 % of its 262
original activity at 1.9 and 2.9 M NaCl after 48 h incubation, respectively. It also retained 22 % and 68 % of 263
its original activity at pH 7 and 12 after 48 h incubation, respectively, and 54 % of its original activity even at 264
100°C after 6 h incubation. Furthermore, the enzyme retained 90–95 % of its activity in 0.5 % (w/v) of the 265
surfactants Tween 40, Tween 60, Tween 80 and cholic acid for 6–48 h incubation, 42–84 % of its activity in 266
the presence of the commercial detergents (0.1 %, w/v) such as rin, surf, ariel, and tide after 7–90 days 267
incubation at 1.89 M Na+ and pH 11, and 75–100 % of its activity in 0.2–1.2 % (w/v) of the oxidizing agents 268
H2O2 and NaClO3. The haloalkaliphilic α-amylase’s amazing stability can be used in applications involving 269
starch liquefaction and the manufacture of detergents and pharmaceutical industries, where high and 270
inhibitory levels of salt, surfactants and detergents are encountered. 271
4.3. Cellulase and xylanase 272
Cellulase and xylanase are enzymes that catalyze the hydrolysis of cellulose and hemicellulose, 273
respectively. They are used as laundry detergent additives and to hydrolyze ligno-cellulosic material in the 274
paper mill industry. So far, a report on cellulase purified from the first anaerobic haloalkaliphilic cellulolytic 275
bacterium Clostridium alkalicellulosi DSM 17461T (formerly designated Clostridium alkalicellum) is the 276
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only instance of haloalkaliphile-derived cellulase (Zvereva et al., 2006; Zhilina et al. 2005). C. alkalicellulosi 277
secreted cellulases with molecular masses of 75 and 84 kDa and high affinity for amorphous or crystalline 278
cellulose. Optimal activity of this cellulase for both microcrystalline and phosphoric acid-swollen cellulose 279
substrate occurred within the pH range 6–9, with more than 70 % of its original activity being retained at pH 280
9.2. The cellulase activity of strain Z-7026T was similar to those of individual clostridial cellulase, but 281
significantly lower than those of intact cellulosome of Clostridium thermocellum. Comparing to the majority 282
of fungal cellulases, it retained more than 50 % of cellulase activity towards microcrystalline cellulose MCC 283
at pH 5–10 and more than 75 % of activity towards phosphoric acid-swollen cellulose at pH 5–9.4. Therefore, 284
this alkaline cellulase offers various opportunities for practical applications (i.e. detergents) where possess the 285
high pH values. Since xylan is more soluble at alkali than neutral conditions, isolation of xylan-degrading 286
microbe and their enzymes from alkaline pH environments has become a greater priority. An alkalistable 287
xylanase was produced by the haloalkalophilic bacterium Staphylococcus sp. strain SG-13, isolated originally 288
from an alkaline soil sample, in wheat bran medium (Gupta et al. 2000). The purified xylanase activity of 289
strain SG-13 has dual pH optima of 7.5 and 9.2 and an optimum temperature of 50°C. This xylanase 290
exhibited a substrate binding capacity of 92 % for hardwood oatspelt xylan but no significant substrate 291
binding for softwood birchwood xylan, and such specificity could be attributed to structural differences of 292
xylan polymers. A promising and cost-effective process for the large-scale production of a halo-alkaline 293
xylanase by the haloalkalitolerant Bacillus pumilus GESF-1 using wheat straw as a carbon source was 294
demonstrated (Menon et al. 2010). Xylanase activity was increased up to 120 % in 1.28 M NaCl, respectively. 295
Approximately 87 and 73 % of original activity was retained in 1.71 and 2.56 M NaCl, respectively, and 296
30 % of original activity was retained at pH 10, respectively. In our laboratory, an anaerobic and xylanolytic 297
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bacterium, designed Alkalitalea saponilacus SC/BZ-SP2T, was derived from Soap Lake, WA (Zhao and Chen, 298
2012). This organism grew at total Na+ concentration of 0.35–1.38 M (optimum at 0.69 M), pH 7.5–10.5 299
(optimum at pH 9.7), and temperature of 8–40 ºC (optimum at 35–37 ºC). It utilized xylan, but no cellulose, 300
from beech wood, birch wood, and oat spelt wood as the sole carbon and energy source under high salinity 301
and alkaline pH, respectively. This implies the secreted extracellular xylanase by strain SC/BZ-SP2 into the 302
growth medium which can be easily purified and used in the paper mill industry for xylan removal. 303
5. Biodegradation and biotransformation 304
Haloalkaliphiles have gained much consideration to treat biological waste treatment and toxic residues 305
from halo-alkaline industrial processes via biodegradation/biotransformation. These compounds include 306
hydrosulfide (HS¯) and sulfide (S2–
), nitriles, benzene, toluene, ethylbenzene and xylenes (BTEX), benzoate, 307
salicylate, and phenol. Among them, HS¯ and S2–
typically are the most dominant sulfur compounds of spent 308
caustics originating from petroleum refineries, which are characterized by sodium concentrations of 309
0.86–2.05 M and a pH value of 9 and greater (Alnaizy 2008, De Graaff et al. 2011). In such combined 310
extreme environments, the haloalkaliphilic sulfur-oxidizing bacteria (SOB) can completely oxidize complex 311
sulfide wastes to sulfate or partially oxidize them to elemental sulfur under low oxygen conditions. In 312
addition, haloalkaliphilic sulfate-reducing bacteria (SRB) can partially oxidize complex sulfide wastes to 313
elemental sulfur though reductive reactions. Partial sulfide oxidation to elemental sulfur under oxygen 314
limitation is more advantageous than complete oxidation to sulfate because it results in regeneration of 315
hydroxyl ions other than protons, thereby limiting the need for caustic absorbent, facilitating separation of 316
elemental sulfur from the final oxidation product, and enabling recirculation of the liquid phase (Sorokin et al. 317
2008b). Many of the sulfide-oxidizing bacteria (SOB) have not yet to be isolated and characterized, but those 318
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belong to four genera of Gammaproteobacteria (i.e. Thioalkalimicrobium, Thioalkalispira, Thioalkalibacter, 319
and Thioalkalivibrio) (Sorokin et al. 2013). De Graaff et al. (2011) reported successful on-site biotreatment of 320
spent caustics from petroleum refineries as an alternative to the physico-chemical treatment using chemical 321
oxidation. For this, sulfide was completely biotransformed into sulfate by a haloalkaliphilic SOB belonging 322
to the genus Thioalkalivibrio at 0.8 M Na+ and pH 9.5 in a continuously fed system with sulfide loading rates 323
of 27 m mol L¯1 day¯
1 and a hydraulic retention time (HRT) of 3.5 days. Similarly, a haloalkaliphilic 324
sulfur-oxidizing mixed culture enriched from Xochimilco alkaline soils (Mexico) containing Thioalkalivibrio 325
and Halomonas strains was active from pH 8–11.5 with a maximum activity at pH 10.6 (Olguín-Lora et al. 326
2011). Clearly, these strains of the genus Thiolalkalivibrio offer great potential for efficient desulfurization in 327
industrial processes at extremely halo-alkaline conditions. A commercial Thiopaq biotechnological 328
application for oxidizing sulfide to elemental sulfur was successfully demonstrated in lab-scale bioreactors 329
inoculated by mix sediments from hypersaline soda lakes (Sorokin et al. 2008b), most of the extremely 330
haloalkaliphilic SOB belongs to the genus Thioalkalivibrio. Fifteen haloalkaliphilic sulfate-reducing bacteria 331
(SRB) were isolated from a soda lake and determined to belong to the genus Desulfonatronum and the genus 332
Desulfonatronovibrio (Sorokin et al. 2011). 333
Compounds containing a C ≡ N (nitrile) are important intermediates in various industrial processes and 334
as building blocks in stereo-selective organic synthesis. They are also natural products formed by cyanide by 335
cyanogenic plants or amino acids by anaerobes (Sorokin et al. 2007a, Sorokin et al. 2008c). Thiocyanate 336
(CNS¯) is produced as a natural product via biological cyanide detoxification processes and as a waste 337
product in coke and metal plants, at high salinity and alkalinity (Sorokin et al. 2001). Several haloalkaliphilic 338
bacteria, Halomonas nitrilicus, Marinospirillum sp., Bacillus alkalinitrilicus, Natronocella acetinitrilica, and 339
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Nitriliruptor alkaliphilus, which used aceto-, propio-, butyro-, iso-butyro-, and valero-nitriles as carbon, 340
energy, and nitrogen sources via nitrile hydratase/amidase pathway, have been isolated (Sorokin et al. 2007a, 341
Sorokin et al. 2007b, Sorokin et al. 2008c). Additionally, several thiocyanate-oxidizing bacteria belonging to 342
SOB genus Thioalkalivibrio were retrieved using on thiocyanate (CNS¯) as electron donor (Sorokin et al. 343
2001, Sorokin et al. 2010). The haloalkaliphilic bacterium Thialkalivibrio thiocyanodenitrificans ARhD 1T 344
could anaerobically oxidized thiocyanate with nitrate as electric acceptor via the identified intermediate 345
cyanate (up to 0.45 mM) and N2O (0.1 mM) to finally form ammonia (65–75% of the metabolized SCN¯ 346
nitrogen) and sulfate (90–95% of the metabolized SCN¯ sulfur) in fed-batch culture at a total Na+ 347
concentration of 0.58 M and pH 9.6 (Sorokin et al. 2004). The halotolerant alkaliphilies Thioalkalivibrio 348
thiocyanoxidans ARh 2T were able to use and degrade thiocyanate as the sole energy and nitrogen source 349
with cyanate as a major intermediate. This organism could also oxidize sulfide, polysulfide, sulfur, and 350
tetrathionate (Sorokin et al. 2002). 351
A variety of toxic aromatic compounds, including BTEX (benzene, toluene, ethylbenzene, and xylenes), 352
benzoate, salicylate, and phenol, are present in petroleum and spent caustic, saline, and alkaline waste 353
streams from refineries (De Graaff et al. 2011). In addition, benzoate is present in food and dye-processing 354
effluents, and salicylate is a key intermediate formed during biodegradation of several polycyclic aromatic 355
hydrocarbons, such as naphthalene and phenanthrene. Combined, the aforementioned aromatic hydrocarbon 356
compounds account for approximately 65 % of bulk-scale chemical waste, are on the priority pollutants list 357
of Environmental Protection Agency (EPA), and can therefore be regarded as models for understanding their 358
biodegradation at the extreme salinity and alkalinity of industrial effluents. Li et al. (2006) first observed that 359
the biodegradations of benzene and its derivatives could simultaneously occur under dual extremes of high 360
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salinity and alkalinity. An aerobic haloalkaliphilie Planococcus sp. strain ZD22, isolated from saline and 361
alkaline soils of the Daqing oil field in China, used benzene as a sole source of carbon and energy. It 362
completely degraded 2 mM benzene at NaCl concentrations of 0.86–3.42 M for 3 days with an optimum of 363
1.71 M, and at pH 7.5–11 with an optimum of pH 9.5. This organism degraded and used not only benzene, 364
toluene, ethylbenzene and o-xylene, but also chlorobenzene, bromobenzene, iodobenzene, and fluorobenzene. 365
Under halo-alkaline conditions (i.e. 0.8 M Na+ and pH 9.5), approximately 93 % of benzene in refinery spent 366
caustics could be removed by a mixed-culture microbial community belonging to the genera Marinobacter, 367
Halomonas and Idiomarina (De Graaff et al (2011). In aerobic batch experiments at salinity of 0.86, 1.50, and 368
1.71 M NaCl and pH 9, Halomonas campisalis 4AT used and degraded both benzoate and salicylate at 369
concentrations of 380 mg/L as carbon and energy sources via the ortho-degradation pathway, as 370
intermetabolites like catechol and cis, cis-muconate were detected (Oie et al. 2007). Three 371
2,4-dichlorophenoxyacetic acid (2,4-D)-degrading bacterial isolates were revived from the highly saline and 372
alkaline Alkali Lake site in Oregon previously contaminated with 2,4-D production wastes (Maltseva et al. 373
1996). The most active of them, strain 1-18 belonging to the family Halomonadaceae , grew optimally on 374
2,4-D at Na+ concentrations of 0.6–1.0 M and pH 8.4–9.4 and degraded up to 3000 mg 2,4-D in 3 days. Strain 375
1-18 used the modified artho-cleavage pathway for 2,4-D degradation, as catechol 1,2-dioxygenase, 376
muconate cycloisomerase, and dienelactone hydrolase were detected. A haloalkaliphilic Nocardioides sp. 377
strain M6 was also isolated from a closed site near Alkali Lake in Oregon that was previously contaminated 378
by high levels of chloro-aromatic compounds (Maltseva and Oriel 1997). It was capable of using 379
2,4,6-trichlorophenol, 2,4-dichlorophenol and 2,4,5-trichlorophenol. These investigations will lead to 380
improved haloalkaliphile biotechnological applications for industrial wastewater treatment in high salt and 381
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high pH systems. 382
Biological denitrification of nitrate-containing waste, such as that found in agricultural run-off and 383
industrial downstream purification steps, is an essential microbially-facilitated process involving a series of 384
intermediate oxide products (NO2–, NO
–, and N2O) of nitrate to ultimately generate gaseous nitrogen at high 385
halolakaline, anoxic, natural / artificial environments. For example, the solution resulting from the 386
replacement of nitrate ions with chloride after industrial ion exchange separation may contain nitrate 387
concentrations of up to 1 g/L at both salt concentration of 0.5–2.0 M and pH of 8–9 (Peyton et al. 2001). 388
Fourteen strains of haloalkaliphilic denitrifiers belonging to genus Halomonas were isolated from extremely 389
saline soda lakes and soils (Shapovalova et al. 2008). All isolates could anaerobically grow on nitrate and 390
nitrite, and in some cases, N2O in saturated sodium carbonate brines (4 M total Na+) and pH 10. When 391
Halomonas sp. AGD 3 anaerobically grew on nitrate at high salinity, high levels of nitrite accumulation 392
occurred. In contrast, no significant level of nitrite and N2O was observed at below 1 M total Na+, indicating 393
that nitrite reduction was a limiting step in the hypersaline denitrification process. A highly active and 394
cytochrome c nitrite reductase (ccNiR) was purified from the haloalkaliphilic Thioalkalivibrio nitratireducens 395
ALEN 2T (Sorokin et al. 2003) molecularly characterized, and could both reduce nitrite and hydroxylamine to 396
ammonia without production of any intermediates and catalyze reductive conversion of sulfite to sulfide 397
(Tikhonova et al. 2006). As a stable hexamer in solution with molecular mass of about 360 kDa, it exhibited a 398
significantly high maximal specific activity (4080 ± 240 µmol/min per mg of the protein) with a reaction rate 399
half-saturation constant Km corresponding to16.7±4.0 mM of nitrite. Recombinant expression and large-scale 400
production of this enzyme is a possibility for the biotechnology industry. Two obligately haloalkaliphilic 401
denitrifiers, Halomonas mongoliensis Z-7009T and Halomonas kenyensis AIR-2
T, were isolated from soda 402
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lakes and capable of reducing nitrate nitrite or N2O as well as oxidizing sulfide (Boltyanskaya et al. 2007). 403
Using acetate, lactate, and glycerol electron donors, Halomonas campisalis 4AT completely reduced 0.5 mg/L 404
nitrate for 3 days at 2.14 M Na+ and pH 9 (Mormile et al. 1999; Peyton et al. 2001). 405
With increasing industrial activity, toxic heavy metal wastes have led to soil and groundwater 406
contamination throughout the globe. Additionally, the radioactive wastes from the experimental detonations 407
of nuclear weapons, the nuclear fuel cycle reprocessing in utility plants, and the medical use of isotopes 408
presents additional problems. The environmental toxicity of some heavy metals at neutral pH conditions can 409
be reduced via their insolubility at alkaline pH conditions. An anaerobic and extremely haloalkaliphilic 410
bacterium strain SLAS-1 was isolated from sediment where it grew via arsenate [As(V)] respiration, using 411
either lactate or sulfide as its electron donor (Oremland et al. 2005). It could reduce arsenate [As(V)] to 412
arsenite As(III) by oxidizing lactate to acetate and HCO3– at optimal salinity range of 20 g/l–saturation and 413
pH 9.5. Several pertechnetate [Tc(VII)O4–]-reducing haloalkaliphiliic Halomonas spp. isolated from 414
soda-lake environments can reduce Tc(VII)O4– to Tc(V), Tc(IV), and Tc(III) (Khijniak et al. 2003). Under 415
anaerobic conditions at 1.05 M Na+ and pH 10, an average of 62 % of the 0.25 mM pertechnetate was 416
reduced to Tc (IV) and Tc(V) by haloalkaliphiles after a 3 days incubation. The isolate SL1 belonging to 417
Halomonas genus from Soap Lake (WA) could reduce Cr(VI) to Cr(III) ion after a 25 days incubation, 418
resulting in the transformation of 75 % of the 0.1 mM Cr(VI) initially present. The maximum specific 419
reduction rate reached 1.6 ± 0.24×10-4
mM Cr(VI) day-1
mg-protein-1
(VanEngelen et al. 2008). 420
6. Biofuel industry 421
Much research on developing biofuels from renewable biomass has been motivated by the rising price of 422
petroleum, recognition of its ultimate depletion, and concerns over global climate change. Halopalkalihiles 423
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have the potential to be applied in the production of biofuel products (e.g. hydrogen, ethanol and carboxylic 424
acids) and other biotechnological products using lignocellulosic biomass as feedstock. For instance, alkaline 425
pretreatment for lignin removal and partial neutralization may create a saline-alkaline environment that is 426
more easily tolerated by haloalkaliphiles during subsequent hydrolysis. In our laboratory, the fermentation 427
products from xylan of beech wood were quantified using 1 % (w/v) by SC/BZ-SP2T (Zhao and Chen, 2012). 428
The identified main end products in the stationary phase were propionate (29.5 mM), acetate (21.2 mM), and 429
minor amounts of burytate (1.51 mM), iso-valerate (0.41 mM) and ethanol (1.21 mM). The pulpmill 430
wastewater from wheat straw generated by alkaline extraction processes is comprised of large quantities of 431
lignin, mono- and di-aromatic compounds, and polysaccharides from cellulose and hemicellulose and very 432
high pH value. Four haloalkaliphilic bacteria of the genus Halomonas were isolated from pulpmill 433
wastewater with a high pH value of 11 (Yang et al. 2008). Among them, Halomonas sp. 19-A and Halomonas 434
sp. Y2 were able to use guaiacol, vanillin, dibenzo-p-dioxin, biphenyl and fluorene at pH 9.5 and 1.71 M 435
NaCl (Yang et al. 2008). These Halomonas species actively growing under extremely halo-alkaline 436
conditions make them promising agents for industrial processes. As a storage and transportation medium for 437
clean and renewable energy, hydrogen is playing an important role in a future energy economy. Elias (2011) 438
filed a patent for haloalkaliphile Halanaerobium hydrogenoformans (formerly designed H. sapolanicus) 439
isolated from Soap lake (WA). This bacterium produced hydrogen, acetate and formate as major metabolic 440
end-products using variety of C-5 and C-6 sugars derived from hemicellulose and cellulose, and grows 441
optimally at 1.2 M Na+ and pH 11 (Begemann et al. 2012; Brown et al. 2011). It can utilize swichgrass and 442
straw that pretreated at low temperature (RT and 55 ºC) than previous report to yield hydrogen with high 443
production rate (0.37 and 0.88 µmol H2/h/ml) in batch reactor. Therefore, it would be expected to offer large 444
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hydrogen production from renewable biomass resources. 445
7. Conclusions and future perspectives 446
Haloalkaliphiles are an interesting and unique class of dual-extremophilic microorganisms thriving 447
under the combined harsh conditions of hyper salinity and extreme alkalinity. They represent the basis for 448
many new and potentially transformative biotechnological efforts aiming to provide novel enzymes and 449
compounds to meet rapidly growing industrial demands for numerous applications. The steady increase in the 450
isolation and identification of novel haloalkaliphiles by the scientific community is accelerating these efforts. 451
The most representative haloalkaliphilies were reviewed, focusing the discussion on their: commercial 452
source for compatible solutes, secondary metabolites, exoezymes, and potential for bioremediation and 453
biotransformation as well as biofuel. Although major advances have been made in the past three decades, 454
compared to the other extremophiles such as halophiles, alkaliphiles and thermophiles, our knowledge of 455
haloalkaliphiles and their exploitable physiology, metabolism, enzymology, and genetics is to date still 456
limited. This is partly due to the high salinity and alkalinity that require specialized media formulations and 457
non-corrodible equipment for large-scale haloalkaliphilic cultivation and subsequent downstream processing 458
for purification. Consequently, conventional industrial efforts for haloalkaliphiles to meet growing market 459
demand are currently hindered. In addition, molecular engineering breakthroughs to generate recombinant 460
mesophilic producers have not been very successful. 461
Further efforts are necessary to improve cultivation biomass concentrations, yields, and volumetric 462
productivities. These will greatly depend on the ability to redirect metabolic fluxes towards the production of 463
targeted products to optimize growth under potentially rigorous conditions. New breakthroughs will not only 464
aid in the discovery / understanding of haloalkaliphiles and enable their commercialization for a multitude of 465
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applications but also allow for their use as sources of ideas for non-haloalkaliphiles. Recent biotechnological 466
advances involving high-throughput enzyme screening, protein engineering, genome sequencing, proteomics, 467
metabolomics, and cloning and heterologous expression in more easily cultivated mesophilic hosts will lead 468
to more novel haloalkaliphilic-based biotechnological applications with exciting new properties. 469
Acknowledgements 470
This paper was supported by the National Natural Science Foundation of China (no. 31370158) and the 471
Basic Research Fund of Chinese Academy of Agricultural Sciences (no. 0042014011). 472
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Ruiz, D.M., Iannuci, N.B., Cascone, O., and De Castro, R.E. 2010 Peptide synthesis catalysed by a 615
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Simon-Colin, C., Raguénès, G., Cozien, J., and Guezennec, J. 2008 Halomonas profundus sp. nov., a new 626
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Table 1. The potentially biotechnological applications of haloalkaliphilic microorganisms in this review. 709
Table 2. The exoenzymes from haloalkaliphiles in this review. 710
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Table 1 711
Microorganism Na+ conc. (M)
growth range (opt.)
pH
growth range (opt.)
Temp. °C
growth range (opt.)
Potential application Reference
Methylophaga alcalica M39T nd– 1.71 (0.51–0.68) 7– 11 (9–9.5) 4– 35 (25–29) ectoine, glutamate Doronina et al. 2003b
Methylophaga lonarensis MPLT 0.01– 1.54 (0.09–0.34) 7– 10 (9–10) 20– 37 (28–30) ectoine, glutamate Antony et al., 2012
Methylophaga murata Kr3T 0.05– 3.0 (0.5–1.5) 6– 10 (8–9) 0– 42 (20–32) ectoine, glutamate Doronina et al. 2005
Methylophaga natronica Bur2T nd– 1.71 (0.34–0.51) 7– 11 (8.5–9) 4– 37 (25–29) ectoine, glutamate Doronina et al. 2003a
Thioalkalimicrobium aerophilum AL 3T 1.2– 1.5 (0.3–0.5) 7.5– 10.6 (9.5–10) nd– 41 (30) ectoine, glutamate Banciu et al., 2005, Robertson et al., 2001
Desulfonatronospira thiodismutans ASO3-1T 1.5– 4.0 (2.0–2.5) 8.5– 10.6 (9.5–10) nd– 43 (35) glycine betaine Sorokin et al., 2011
Natranaerobius thermophilus JW/NM-WN-LFT 3.1– 4.9 (3.3–3.9) 8.3– 10.6 (9.5) 35– 56 (53) glycine betaine Zhao et al., (Data not published)
Thioalkalivibrio versutus ALJ 15T nd– 4.3 (1.0–2.0) 7.5– 10.6 (10) nd– 47 (35) glycine betaine Banciu et al., 2005; Robertson et al., 2001
Natronomonas pharaonis SP1T 2.0– 5.1 (3.5) 8– 11 (8.5–9) nd– 53 (43–45) halorhodopsin (hR); phoborhodopsin (ppR) Duschl et al. 1990; Hirayama et al. 1992;
Natronomonas pharaonis DSM 2160T 2.0– 5.1 (3.42) 8– 11 (8.5–9) nd– 53 (45) halorhodopsin (hR) Ihara et al., 2008; Xu et al. 1999
Halomonas profundus AT1214T 0.34– 0.51 (nd) 8–9 (nd) 32– 37 (nd) poly-3-hydroxybutyrate-co-3-hydroxyvalerate (PHV);
poly-3-hydroxybutyrate (PHB)
Simon-Colin et al. 2008
Halomonas campaniensis 5AGT 0– nd (1.71) 7– 10 (9) 10– 43 (37) poly-3-hydroxybutyrate (PHB) Gambacorta et al. 2005; Strazzullo et al. 2008
Halomonas campaniensis MCM B-1027 0– 4.0 (1.0) 7– 11 (9) 4– 45 (30) hydroxybutyrate-co-hydroxyvalerate
(PHB-co-PHV) copolymer
Kanekar et al. 2011; Kulkarni et al. 2010
Methylophaga murata Kr3T 0.05– 3.0 (0.5–1.5) 6– 10 (8–9) 0– 42 (20–32) exopolysaccharide (EPS) Doronina et al. 2005
Bacillus sp. Vel 0–3.5 (1.7) 7.0–9 (8–8.5) 37 (nd) protease Patel et al. 2005; Patel et al. 2006
Bacillus sp.AH-6 nd 10 (nd) 37 (nd) protease Dodia et al. 2008a, 2008b
Salinivibrio costicola 18AGT 0.1–4.3 (2.0) 7–10.5 (9) 10–40 (30) protease Lama et al. 2005, Romano et al. 2005
Geomicrobium sp. EMB2 0.85–3.42 (2.05) 7–10 (8.5) 30 (nd) protease Karan et al
Natrialba magadii ATCC 43099T 2.0–5.1 (4.1) 8.5–11.5(9.5) 37 (nd) protease Tindall et al. 1980; Mwatha & Grant. 1993
Natrialba pharaonis DSM 2160T 4.0 (nd) 9.5 (nd) 37 (nd) protease Stan-Lotter et al. 1999
Natrialba occultus NCBM 2192T 3.59 (nd) 10 (nd) 37 (nd) protease Studdert et al. 1997, 2001
Natronococcus sp. Ah-36 1.37–5.13 (2.56–3.42) 8–10 (9) 20–55 (40–45) α-amylase Kobayashi et al. 1992
Chromohalobacter sp. TVSP 101 0.85–5.13 (3.42) 6–10 (9) 35–55 (37) α-amylase Prakash et al. 2009
Saccharopolyspora sp. A9 4 (nd) 11 (nd) 37–55 (55) α-amylase Chakraborty et al. 2011
Clostridium alkalicellulosi DSM 17461T 0.02–0.4 (0.15–0.3) 8–10.2 (9) 18–42 (35–40) cellulose, xylanase Zhilina et al. 2005; Zvereva et al. 2006
Staphylococcus sp. SG-13 0.43–2.56 (nd) 6–11 (8) 25–50 (37) xylanase Gupta et al. 2000
Bacillus pumilus GESF-1 0.85–2.56 (0.85) 8–10 (8) 37 (nd) xylanase Menon et al. 2010
nd, not determined in the publications.712
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Continue Table 1 713
Microorganism Na+ concentration (M)
growth range (opt.)
pH
growth range (opt.)
Temp. °C
growth range (opt.)
Potential application Reference
Thioalkalivibrio spp. 0.8 (nd) 9.5 (nd) 35 (nd) sulfur-oxidizing De Graaff et al. 2011
Thioalkalivibrio spp. 0.8 (nd) 8– 11.5 (10.6) 35 (nd) sulfur-oxidizing Olguín-Lora et al. 2011
Halomonas spp. 0.6– 1.7 (nd) 7– 11 (9.8) 30 (nd) sulfur-oxidizing Olguín-Lora et al. 2011
Thioalkalivibrio spp. 2– 3 (nd) 10 (nd) 30 (nd) sulfid → elemental sulfur Sorokin et al. 2008b
Desulfonatronum spp. 0.2– 2.5 (0.5–1) 8.0– 10.4 (9.3–9.5) 30 (nd) sulfate-reducing Sorokin et al. 2011
Desulfonatronovibrio spp. 0.2– 1.75 (0.5) 8.5– 10.3 (9.5–10) 30 (nd) sulfate-reducing Sorokin et al. 2011
Thialkalivibrio thiocyanodenitrificans ARhD 1T 0.3– 1.8 (nd) 9.6– 10 (nd) 30 (nd) biodegradation of thiocyanate Sorokin et al. 2004
Thioalkalivibrio thiocyanoxidans ARh 2T 0.3– 4 .3 (0.5–1) 8.5– 10.5 (10.2) 30 (nd) biodegradation of thiocyanate; sulfur-oxidizing Sorokin et al. 2002
Planococcus sp. ZD22 0.08– 4.27 (1.71) 7.5– 11 (9.5) 2– 36 (20–25) biodegradation of benzene Li et al. 2006
Halomonadaceae sp. 1-18 0.6– 1 (nd) 6.5– 10 (9–9.5) biodegradation of 2,4-dichlorophenoxyacetic acid Maltseva et al. 1996
Nocardioides sp. M6 0– 1.35 (0.2–0.4) 8.8– 9.8 (9.2) 30 (nd) biodegradation of 2,4,6-trichlorophenol,
2,4-dichlorophenol, and 2,4,5-trichlorophenol
Maltseva and Oriel 1997
Halomonas spp. 4 (nd) 10 (nd) 30 (nd) denitrification Shapovalova et al. 2008
Thioalkalivibrio nitratireducens ALEN 2T 0.2– 1.5 (0.4–0.5) 8.5– 10 (9.5–10) 35 (nd) denitrification Sorokin et al. 2003
Halomonas mongoliensis Z-7009T 0.16– 2.2 (0.7–1.7) 8– 10.5 (8.5–9.6) 15– 50 (36–40) oxidization of sulfide and reduction of nitrous oxide Boltyanskaya et al. 2007
Halomonas kenyensis AIR-2T 0.04– 2.2 (0.5–1.2) 7.5– 10.6 (9.5) 15– 55 (36–40) oxidization of sulfide and reduction of nitrate Boltyanskaya et al. 2007
Halomonas campisalis 4AT 0.2– 4.5 (1.5) 6– 12 (9.5) 4– 50 (30) biodegradation of benzoate and salicylate,
reduction of nitrate
Mormile et al. 1999; Peyton et al. 2001;
Oie et al. 2007
Halanaerobacteriales sp. SLAS-1 3.42– 5.64 (4.27–5.64) 9– 10.5 (9.5) nd (30) arsenate [As(V)] → arsenite [As(III)] Oremland et al. 2005
Halomonas spp. 1.05 (nd) 10 (nd) 30 (nd) Tc(VII)O4– → Tc(V), Tc(IV) and Tc(III) Khijniak et al. 2003
Alkalitalea saponilacusSC/BZ-SP2T 0.35– 1.38 (0.44–0.69) 7.5– 10.5 (9.7) 8– 40 (35–37) production of propionate and acetate Zhao and Chen 2012
Halanaerobium hydrogenoformans ATCC PTA-10410T 0.81– 2.94 (1.58) 7.5– 12 (11) 33 (nd) production of biohydrogen Begemann et al. 2012; Brown et al. 2011
Elias et al. 2011
nd, not determined in the publications. 714
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Table 2 715
Exoenzymes from haloalkaliphiles Na+ concentration (M)
range (opt.)
pH
range (opt.)
Temp.°C
range (opt.) Reference
Protease from Bacillus sp. Vel 0– 0.17 (0.03) 8.5– 12.0 (10–11) 5– 50 (37) Gupta et al. 2005
protease from Bacillus sp.AH-6 0– 4 (0.15–0.2) 8– 13 (9–11) 37– 80 (50) Dodia et al. 2008a, 2008b
Protease from Salinivibrio costicola 18AGT 0– 2.57 (0.34) 5– 12 (8) 5– 100 (50) Lama et al. 2005
Protease from Geomicrobium sp. EMB2 0– 3.42 (0.85) 6– 12 (10) 0– 70 (50) Karan et al. 2011
Protease from Natrialba magadii ATCC 43099T 0.5– 2 (1–1.5) 6– 12 (8–10) 0– 70 (60) Giménez et al. 2000
Protease from Natrialba pharaonis DSM 2160T 0– 4 (0.5–4) 6– 12 (10) 0– 70 (61) Stan-Lotter et al. 1999
Protease from Natrialba occultus NCBM 2192T
0.1– 4 (1–2) 5.5– 12 (7–9) 30– 70 (60) Studdert et al. 1997, 2001
α-amylase from Natronococcus sp. Ah-36 1– 5(2.5) 4.2– 10.5 (8.7) 30– 60 (55) Kobayashi et al. 1992
α-amylase from Chromohalobacter sp. TVSP 101 0– 5.12(0–3.41) 5– 10 (9) 30– 80 (65) Prakash et al. 2009
α-amylase from Saccharopolyspora sp. A9 0.60– 2.9(1.88) 4– 12 (10–12) 55– 100 (55–95) Chakraborty et al. 2011
Cellulase from Clostridium alkalicellulosi DSM 17461T nd 4– 11 (6–9) nd Zvereva et al. 2006
Pylanase from Staphylococcus sp. SG-13 nd 6.0– 10.5 (7.5–9.2) 10– 65 (50) Gupta et al. 2000
Xylanase from Bacillus pumilus GESF-1 0– 2.56 (0.85) 7– 13 (7–8) 30– 70 (40) Menon et al. 2010
nd, not determined in the publications 716
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