Bioresource Technology 153 (2014) 189–197

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Bioresource Technology

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Production and characterization of biodiesel from carbon dioxideconcentrating chemolithotrophic bacteria, Serratia sp. ISTD04

0960-8524/$ - see front matter � 2013 Elsevier Ltd. All rights reserved.http://dx.doi.org/10.1016/j.biortech.2013.11.075

⇑ Corresponding author. Tel.: +91 11 26704321 (O), +91 11 26191370 (R); fax:+91 11 26717586.

E-mail addresses: isthakur@hotmail.com, isthakur@mail.jnu.ac.in (I.S. Thakur).

Randhir K. Bharti, Shaili Srivastava, Indu Shekhar Thakur ⇑School of Environmental Sciences, Jawaharlal Nehru University, New Delhi 110 067, India

h i g h l i g h t s

�Microbial community isolated from marble rock for CO2 sequestration.� Bacterium indicated presence of carbonic anhydrase and RuBisCO activity.� Western blot analysis confirmed RuBisCO activity in bacterium.� Hydrocarbon and lipid extracted from cell lysate of Serratia sp.� Transesterification indicated formation of biodiesel from lipids.

a r t i c l e i n f o

Article history:Received 14 October 2013Received in revised form 15 November 2013Accepted 25 November 2013Available online 6 December 2013

Keywords:BiodieselCarbonic anhydraseChemolithotrophic bacteriumCarbon dioxide sequestrationRuBisCO

a b s t r a c t

A chemolithotrophic bacterium, Serratia sp. ISTD04, enriched in the chemostat in presence of sodiumbicarbonate as sole carbon source was evaluated for potential of carbon dioxide (CO2) sequestrationand biofuel production. CO2 sequestration efficiency of the bacterium was determined by enzymaticactivity of carbonic anhydrase and ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO). Further,Western blot analysis confirmed presence of RuBisCO. The bacterium produced 0.487 and 0.647 mg mg�1

per unit cell dry weight of hydrocarbons and lipids respectively. The hydrocarbons were within the rangeof C13–C24 making it equivalent to light oil. GC–MS analysis of lipids produced by the bacterium indicatedpresence of C15–C20 organic compounds that made it potential source of biodiesel after transesterifica-tion. GC–MS, FTIR and NMR spectroscopic characterization of the fatty acid methyl esters revealed thepresence of 55% and 45% of unsaturated and saturated organic compounds respectively, thus making ita balanced biodiesel composition.

� 2013 Elsevier Ltd. All rights reserved.

1. Introduction

The increasing use of fossil fuels for energy since the beginningof the industrial age has released a tremendous quantity of CO2

into the atmosphere. Producing and using biomass-derived fuelsfor transportation can reduce GHGs emission. Increase in carbondioxide concentration in atmosphere stimulates the autotrophiccarbon fixation of biota that is mediated by carboxylating enzymesand ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCO)(Jo et al., 2013). Photosynthetic organisms convert light energy togenerate reducing equivalents from water, thus allowing fixationand use of carbon dioxide as a growth substrate (Jo et al., 2013).Carbon dioxide and dissolved inorganic carbon (bicarbonate) areessential components for the growth of microorganisms (Kurianet al., 2006). One of the most effective methods to mitigate

challenges of the rising levels of CO2 is sequestration by microor-ganism since some microbes are capable of fixing atmosphericcarbon dioxide into valuable products like calcite, vaterite andaragonite. The ability of certain bacterial strains to precipitatecarbonates has been previously reported from Bacillus cereus,Pseudomonas aeruginosa, Pseudomonas putida (De Muynck et al.,2010), Pseudomonas fluorescens, Bacillus subtilis (Zamarreno et al.,2009), Bacillus pasteurii (Tao and Wenkun, 2012) and Myxococcusxanthus (Rodriguez-Navarro et al., 2003).

The CO2 acquisition mechanism is termed as carbon concentrat-ing mechanism (CCM) and it has been suggested that carbonicanhydrases (CA) plays a key role in bacteria (Dou et al., 2008). Car-bonic anhydrases, CA (EC 4.2.1.1) catalyzes the reversible dehydra-tion of HCO�3 to CO2. This reaction is known to play important rolesin various biological processes such as ion exchange, respiration,pH homeostasis, CO2 acquisition, and photosynthesis (Smith andFerry, 2000). The carbonic anhydrase functions to convert an accu-mulated cytosolic pool of HCO�3 into CO2 within the carboxysome.The generation of CO2 coupled with a diffusive restriction to the

190 R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197

efflux from the carboxysome, possibly imposed by the proteinshell, leads to the localized elevation of CO2 around the active siteof ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO)within the carboxysome. The substrate for the carboxysome,HCO�3 , is accumulated in the cytosol by the operation of a numberof active CO2 and HCO�3 , transporters. These transporters are lo-cated on plasma membrane and exist in both low affinity and highaffinity transporter forms (Murray et al., 2003). The use of biotech-nological approaches for the sequestration of carbon dioxide is theemerging technology and is considered to be environmentally be-nign. Based on above facts microorganisms may be enriched in thechemostat in presence of bicarbonate as sole carbon source to ac-quire proteins and genes for enhanced sequestration processes.Thus, bacteria that contain CA and RuBisCO might be raised as goodcandidates for biotechnological application for CO2 sequestration.

There is very limited information related to production of bio-diesel from bacteria. Fatty acids, alcohols and alka(e)nes are basicbiomolecules for production of biofuel. Biofuel is a generic termfor alternate energy resource produced from renewable biologicalsubstances. They can be a substitute or an extender for the conven-tional petroleum-based diesel. Biodiesel is defined as a fuel con-sisting of mono-alkyl esters of long-chain fatty acids originatingin fats, oils or lipids. Lipids from cynobacteria, algae, jatropha, palmtrees and soybeans have been used to produce biodiesel (Schenket al., 2008). Biomass from microbial photosynthesis is being ex-plored for biofuels production by genetic engineering (Howardaet al., 2013). Bacteria possess significant advantages over photo-synthetic organisms such as higher plants and microalgae, whichhave been the main focus of attention, for biological fuel oil pro-duction. These advantages include a much faster life cycle and ra-pid growth rate, the ability to live in a high-density culture andfurther, microbial photosynthesizers can be genetically modifiedmore easily. Hydrocarbons equivalent to light oil alternativesformed the major components of the lipids produced by bacterium.

Bacteria are intrinsically capable of synthesizing fatty acids,which are precursors in the biosynthesis of their cell envelopes(Moazami et al., 2011). Fatty acids in bacteria are formed from acet-yl-CoA with ATP as the energy source and NADPH as the source ofreducing equivalents, similar to plants. Acetyl-CoA carboxylase(ACC) catalyzes fatty acid biosynthesis through the formation ofmalonyl-CoA from acetyl-CoA and bicarbonate in an ATP-dependentmanner. Fatty acyl-ACPs (acyl carrier proteins) are synthesized frommalonyl-CoA by a multi-subunit fatty acid synthase (FAS). The fattyacyl moiety is eventually transferred onto glycerol derivatives (orother alcohols) by glycerol-3-phosphate acyltransferase and formslipids (Lu et al., 2008; Rottig and Steinbüchel, 2013).

Alkanes are produced directly from fatty acid metabolites bydecarbonylation of fatty aldehydes (Schirmer et al., 2010). Thepathway consists of an acyl–acyl carrier protein reductase and analdehyde decarbonylase, which together convert intermediates offatty acid metabolism to alkanes and alkenes (Schirmer et al.,2010). The synthesis of hydrocarbons by microorganisms dependsconsiderably on the growth conditions that provide a way for itsphysiological regulation. The processes for microbiological produc-tion of extracellular aliphatic and volatile non-methane hydrocar-bons are straight chain-, branched-, volatile non-methanehydrocarbons, and isoprenoids. Mechanisms of the hydrocarbonsynthesis appear to be different in various microorganisms. Thehydrocarbon contents vary among different systematic groups ofplants and microorganisms. The production and secretion of C13–C17 mixtures of alkanes and alkenes are reported in photoautotro-phic bacteria (Schirmer et al., 2010). Hydrocarbons of theappropriate size are ideal fuels, meeting all the previously indi-cated fuel criteria. Gram-positive bacteria of the genus Clostridiumsp., produced intracellular hydrocarbons from C11 to C35 with thepredominance of middle-chain n-alkanes (C18–C27) and long-chain

n-alkanes (C25–C35) (Ladygina et al., 2006). A halotolerant bacte-rium Vibrio furnissii has been reported to produce intracellularand extracellular hydrocarbons (C15–C24) is similar to keroseneand light oil (Park et al., 2005). Yeasts are able to synthesize a widerange of hydrocarbons from C10 to C34, which include not only n-al-kanes but also unsaturated and branched components (Ladyginaet al., 2006). The hydrocarbon profile of fungal spores is similarto that of higher plants and is dominated by the odd-numberedn-alkanes (C27, C29, and C35) (Ladygina et al., 2006).

In this study, bacteria from the marble rock are enriched in thechemostat in presence of sodium bicarbonate as sole carbon sourceso that it can sequester carbon dioxide more efficiently, and in-creased biomass can be exploited for biodiesel production. The car-bon dioxide sequestration capacity of the bacterium was evaluatedby carbonic anhydrase and RuBisCO activity. The inherent ability ofthe chemolithotrophic bacteria of marble mine rock to sequestercarbon dioxide was evaluated in terms of biomass production,hydrocarbon and lipid content and biodiesel quality.

2. Methods

2.1. Sampling site

Sediment samples were collected from marble rocks of the pal-aeoproterozoic metasediments of the Aravali Supergroup, Alwararea (27� 340 N, 76� 380 E), Rajasthan, India, for isolation of mi-crobes. The upper portion of the rock was scrapped carefully fromdifferent places and dissolved in autoclaved distilled water (1:10w/v) which was used as inoculum for enrichment of bacteria inthe chemostat by continuous culture (Thakur, 1995).

2.2. Microorganism and culture conditions

A chemostat culture was set up in a 2-L glass vessel, effectivevolume 1 L, with culture condition as follows–stirring at150 rpm; temperature at 30 �C; and pH 7.6 in the minimal saltmedium (MSM). The composition of MSM (g L�1) was: Na2HPO4,7.8; KH2PO4, 6.8; MgSO4, 0.2 g; NaNO3, 0.085; ZnSO4.7H2O, 0.05;Ca(NO3)2.2H2O, 0.05; FeSO4.7H2O 0.4; Na2S.9H2O, 1.87; (Thakur,1995). The supernatant containing microorganisms (50 mL) ofmarble rock was used as inoculums in the chemostat. The bacteriawere enriched in the chemostat initially in the presence of 20 mMsodium bicarbonate. Then the concentration of sodium bicarbonatewas increased to 50, 100, and 150 mM. Growth of microorganismswas determined each time and after stabilization of growth con-centration of sodium bicarbonate was increased. Bacterial commu-nity was maintained in MSM and 5% gaseous CO2 in shake flasks.The carbon dioxide concentration (5%) was maintained in a carbondioxide incubator (Sanyo, Japan) equipped with an infrared sensor.

Culture medium obtained from different concentration (20, 50,100 and 150 mM) of sodium bicarbonate was diluted serially andthe supernatant (100 lL) was spread on LB-agar plate for evaluationof diversity of bacterial population. The bacterium adapted at higherconcentration of sodium bicarbonate re-cultured on minimal saltmedium having sodium bicarbonate (150 mM). Genomic DNA fromthe bacterial strain was isolated with the Genome DNA Kit (QiagenInc., USA). The 16S rRNA gene was amplified from genomic DNA byusing PCR with universal primers, sequenced and phylogeneticplacement was carried out as described earlier (Jaiswal et al., 2011).

2.3. Culture conditions for assay of carbon dioxide concentratingenzymes

Bacterial strain was grown in minimal salt media andNaHCO3 (50 mM) for a period of 48 h at 30 �C. Culture medium

R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197 191

was centrifuged at 8000 rpm for 10 min. Cell pellet was washedand suspended in sonication buffer (10 mL) containing Tris–HCl(50 mM, pH 7.6), dithithretiol (1 lM), 1 mM PMFS and lysozyme(0.2%). Cells were disrupted through sonication at 20,000 htz for8 min, at 20-s interval at 4 �C. The lysate was centrifuged at14,000 rpm for 30 min and supernatant was taken as crude ly-sate. Total protein concentration was measured in the lysate asdescribed by Bradford using BSA as a standard. The carbonicanhydrase activity was measured in aqueous phase by usingnitrophenyl esters (Innocenti et al., 2008). The enzymatic reac-tion contained Tris–HCl buffer (50 mM, pH 7.5), 3 mM p-nitro-phenyl acetate (p-NPA), enzyme preparations (100 lL). Oneunit of enzyme activity was expressed as 1 lmol of p-nitro-phenyl acetate hydrolyzed per minute. RuBisCO was determinedusing spectrophotometric method (Yeates et al., 2008). RuBP-dependent oxidation of NADH was used to monitor activity ofRuBisCO. In crude lysate and protein fractions, the initial rateof oxidation of NADH was determined by recording the decreasein A340, using a e340 of 6.220 lM�1cm�1. Reactions were per-formed in 1.0-ml quartz cuvettes with a 1-cm light path. Activ-ities were assayed at 4 �C by adding 100 ll of crude protein to500 lL of 200 mM sodium Tris–Cl buffer containing 10 mMMgCl2, 132 mM KHCO3, 10 mM DTT and 5.5 mg ml�1 ATP at pH7.8. Then 100 lL of 5 Unit each of coupling enzymes (GPDHand PGK), 100 lL of 2 mM NADH were added and mixed well,followed by addition of 200 lL of 2.5 mM RuBP. It was mixedquickly and effectively. Activity of RuBisCO was recorded usingspectrophotometer and was expressed as the amount of enzymethat catalyzes the oxidation of 1 lmol of NADH per min. Fourlmoles of NADH are oxidized for each lmole of D-ribulose 1,5-diphosphate utilized. One unit enzyme will convert 1.0 lmol ofD-RuDP and CO2 to 2.0 lmol of D-3-phosphoglycerate per minuteat pH 7.8.

Sodium dodecyl sulfate polyacrylamide gel electrophoresis(SDS–PAGE) of crude lysate was performed. In this method, acryl-amide gel (12%) was used, and electrophoresis was performed.Gels were stained by Comassie brilliant blue as described by(Mishra and Thakur, 2011). Unstained Protein Molecular WeightMarker, a mixture of seven native proteins (14.4–116 kDa) wasused, in electrophoresis (SDS–PAGE). For immunoblot analysis,crude protein of cell lysate separated by 12% SDS–PAGE wastransferred into nitrocellulose membrane with Mini trans-blotelectrophoretic transfer cell (GE Healthcare) according to direc-tions provided by the manufacturer (Ashida et al., 2005). Aftertransferring, membrane was incubated in blocking solution con-taining 5% skimmed milk in 1� TBS for 12 h at 37 �C in a slowrocker. This was followed by washing with 1� TBST twice for20 min and a final rinse with 1� TBS. Membrane was then incu-bated for 4 h at 37 �C with primary antibody [Anti-RuBisCO(plant) antibody produced in chicken] against rbcL (1:500). Thiswas followed by washing with 1� TBST twice for 20 min and afinal rinse with 1� TBS. Membrane was then subjected to treat-ment with peroxidase conjugated goat anti-chicken IgG dilutedto 1:10,000 (secondary antibody) for 45 min. This was again fol-lowed by washing with 1� TBST for 20 min followed by a finalrinse with I� TBS. Horseradish peroxidase-labeled secondaryantibodies and enhanced chemiluminescence (Amersham) wereused for detection of the antibody–antigen conjugate. The anti-body antigen complex was then detected using enhanced chemi-luminescence (ECL). For this, 10 mL of 100 mMTris–Cl (pH 8.5)containing 50 ll of 250 mM luminol, 22 lL of 90 mM p-coumaricacid and 4 lL hydrogen peroxide was added into the membrane.The chemiluminesce was visualized with the help of image quantLAS 4000 imager (GE healthcare) and images were acquired with

software.

2.4. Extraction of bacterial lipids and hydrocarbons

Lipid was extracted from bacterial cell lysate obtained aftersonication and supernatant of bacterial cells. Then cell lysate wasmixed with supernatant. For extraction of lipid, 50 mL lysate andsupernatant was mixed with 100 mL chloroform/methanol (2:1v/v) by vigorous shaking for 30 min. The organic phase (extract)was separated by a separating funnel, and the lower phase contain-ing lipids were finally concentrated on a rotary evaporator, andthen transferred to a pre-weighed glass test tube. For extractionof hydrocarbon, 50 mL lysate and supernatant was mixed with100 mL ethyl acetate by vigorous shaking for 30 min. The organicphase (extract) was separated by a separating funnel, and the ex-tract was finally concentrated on a rotary evaporator. Then it wasre-dissolved in 100 lL acetonitrile. The extract was used for iden-tification of lipids and hydrocarbons by using gas chromatographymass spectroscopy (GC–MS) analyses and FTIR (Fourier transformsinfrared spectroscopy) and NMR (nuclear magnetic resonance)spectroscopy.

2.5. Transesterification processes for production of biodiesel

Experiments were planned to study the effect of differentcatalysts and their concentration, reaction temperature, oil/meth-anol ratio, and stirring intensity on the transesterification of bac-terial lipids. Catalysts used included sodium hydroxide andpotassium hydroxide. The catalyst (NaOH) concentrations takenwere 0.25%, 0.50%, 0.75%, 1.00%, 1.25%, and 1.50%. The reactiontemperatures chosen were 20, 25, 30, 35, 40, and 45 �C. The stir-ring intensity was also varied and included 150, 200, 250, 300and 350 rpm. The time duration opted was 1, 2, 3, and 4 h. Aftershaking, the solution was kept for 16 h to settle the biodiesel andthe sediment layers clearly. After the end of the reaction, thereaction mixture was allowed to cool and equilibrate, resultingin the separation of two phases. The upper phase consisted ofFAMEs, and the lower phase contained glycerol, the excess meth-anol, the remaining catalyst together with the soaps formed dur-ing the reaction, and some methyl esters and partial glycerides.After separation of the two layers, the upper FAME layer waswashed by 5% water. The remaining catalyst was removed bysuccessive rinses with distilled water. Finally, the residual waterwas removed by treatment with Na2SO4, followed by filtrationand evaporation in a rotator evaporator.

Yield of methyl esters = (grams of methyl esters produced/grams of oil taken for the reaction) � 100

After separation and purification of FAMEs/Biodiesel, it was re-dissolved in 1 ml methanol and analyzed by GC–MS, NMR andFTIR.

2.6. Gas chromatography–mass spectroscopy (GC–MS)

The hydrocarbons and lipids were identified by using gaschromatography–mass spectroscopy (GC–MS) (Varian) equippedwith a capillary column (DB-5 MS; 0.25 lm film thickness,0.25 mm i.d., 30 m in length). One lL of each sample was analyzedby GC–MS at conditions: split less mode; initial temperature 80 �Cheld for 2 min; temperature increased from 80 to 200 �C at a rate of10 �C min�1 held for 6 min on reaching 200 �C; temperature fur-ther increased from 200 to 300 �C at a rate of 5 �C min�1 held for15 min on reaching 300 �C. The head pressure of the helium carriergas was 81.7 kPa; helium flow rate 1.21 mL min�1. Data wascompared with the inbuilt standard mass spectra library system(NIST-05 and Wiley-8) of GC–MS.

192 R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197

2.7. Analysis of functional groups by Fourier transforms infraredspectroscopy (FTIR)

The functional groups present in the lipids were identified usingFourier transform infrared spectroscopy (FTIR). For FTIR analysis,lipid sample was mixed with KBr in a ratio of 5:100 to make theKBr discs for spectrum analysis. The analysis was done with thehelp of FTIR spectrometer (model Varian 7000 FTIR) (Schmitt andFlemming, 1998).

2.7.1. 1H NMR and 13C NMR measurements of lipidsNMR experiments were performed at 7.05 T using a Varian Mer-

cury Plus NMR spectrometer equipped with 5 mm Varian probes(ATB and SW) using CDCl3 as solvent (Laetitia et al., 2006). A 1H(300 MHz) spectrum was recorded with pulse duration of 45�, a re-cycle delay of 1.36 s and 16 scans. The spectra were referenced totetramethylsilane (ä) 0.0 ppm). 13C (75.46 MHz) spectra was re-corded with a pulse duration of 45�, a recycle delay of 0.28 s and300 scans. The spectra were referenced to CDCl3 (ä) 77.0 ppm.

3. Result and discussion

3.1. Enrichment of bacterial community for utilization of carbondioxide

Six morphologically different bacterial isolates (A–F) were cul-tured on LB-agar plate from the bacterial community enriched inthe chemostat at 20 mM and 50 mM NaHCO3; however, three iso-lates were obtained at 100 mM, and single isolate at 150 mM. Bac-terial colonies that appeared on agar plates were differentiatedmorphologically by shape, size, color, smoothness, texture, marginetc. Members of community grown on LB-agar were tested for sur-vival pattern by growth curve (data not shown). The result of thestudy showed higher growth of isolate D in comparison to otherfive isolates. There were initial changes in growth and utilizationof sole carbon source (sodium bicarbonate) by the other isolatesbut isolate D had better capability to utilize CO2 obtained fromNaHCO3 as a carbon source. In the experiment, bacterial isolateswere part of indigenous marble rock, which were initially part ofcalcium carbonate environment and enriched in the chemostat.

Time (ho

0 20 40

S

peci

fic a

ctiv

ity o

f enz

yme

carb

onic

anh

ydra

se (µ

mol

e PN

PA /m

in/m

g pr

otei

n)

10

15

20

25

30

35

40

specific activitygrowth of bacteria

Fig. 1. Growth of bacterium, Serratia sp.ISTD04, isolated from bacterial community enricsalt medium with sodium bicarbonate as sole carbon source. At right y2-axis, dotted liney1-axis, bold line represents specific activity of crude enzyme (carbonic anhydrase) wit

The community had dominant chemolithoautotrophs microorgan-isms which initially used sodium bicarbonate as a source of carbonfor their survival and metabolism.

The bacterial isolate (D) showing higher tolerance and growthin presence of sodium bicarbonate (150 mM) was selected foridentification by 16S rDNA sequencing method. The resulting se-quence was entered into the BLAST nucleotide search program ofthe National Center for Biotechnology Information to obtain closelyrelated phylogenetic sequences indicated similarity of the isolateto Serratia sp. Pair wise alignment giving closest match was chosenand phylogeny tree was drawn using MEGA 3.1 software (Jaiswalet al., 2011). Molecular techniques based on 16S rDNA genes haveprovided new insights to elucidate microbial community. Sequenc-ing of 16S rRNA genes has been very useful for describing the com-positions of bacteria (Jaiswal et al., 2011).

3.2. Production and assay of carbon dioxide concentrating bacteria

The activity of carbonic anhydrase was determined by nitro-phenyl esters methods. It is difficult to measure the activity offree or immobilized enzyme with gas phase substrate such asCO2. For this reason, in the present work carbonic anhydraseactivity was measured in aqueous phase by using nitrophenyl es-ters method. Formation of carbonic anhydrase in cell lysate wasdetermined from 0 to 75 h. It was observed that carbonic anhy-drase activity determined after 10 h in cell extract lysate whichreached to maximum at 50 h and then declined (Fig. 1). Resultsof the study indicated that formation of carbonic anhydrase en-zyme was similar to the growth of bacterial cells. Higher activityof carbonic anhydrase in experiment revealed that the bacterialstrain isolated was efficient for CO2 sequestration because ofthe presence of carbonic anhydrase which is a supporting enzymefor the activity of RuBisCO. The enzyme is thought to dehydrateabundant cytosolic bicarbonate and provide ribulose 1,5-bisphos-phate carboxylase/oxygenase (RuBisCO) sequestered CO2 withinthe carboxysome (Dou et al., 2008). Some chemolithotropic bac-teria and most of cyanobacteria have proper channels for trans-port of dissolved inorganic carbon (CO2, HCO�3 , and CO2�

3 ). Byenhancing the bicarbonate ion concentration or gaseous CO2 con-centration in both the cases, bacteria accumulates HCO�3 in

ur)

60 80 100

O.D

at 5

95 n

m

(gr

owth

of b

acte

ria)

0.1

0.2

0.3

0.4

0.5

0.6

0.7

hed in sodium bicarbonate (50 mM) in the chemostat and further grew in minimalrepresents bacterial growth at O.D. 595 nm with respect to time in hours and at left

h respect to time in hour and at x-axis.

R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197 193

cytoplasm and triggers higher CA activity and conversion of HCO�3to CO2 (Ramanan et al., 2009). In these way bacteria gets ac-quainted to sequester more CO2. So activity of carbonic anhydraseis important factor for CO2 sequestration. Carbonic anhydrases(CA) have been associated in fixation of dissolved inorganic car-bon as reported from hydrothermal vent habitat and other eco-systems (Ramanan et al., 2009). Five different classes ofcarbonic anhydrase have been identified so far in which threewere isolated and characterized from bacterial system (Smithand Ferry, 2000). Genes encoding homologous of all three classesof carbonic anhydrases are reported from T. crunogena genome,proteobacteria and chemolithotrophic bacteria (Smith and Ferry,2000). But in this study a noble bacterium and its carbonic anhy-drase from marble rock was isolated and it was characterized soas to use it for sequestration of carbon dioxide.

RuBisCO enzyme was assayed by spectrophotometeric meth-ods. The RuBisCO activity in crude enzyme was determined from0 to 120 h which reached to maximum at 72 h (8.94 nmol CO2 -min�1 mg�1 protein) after that it was declined (Fig. 2a). In this case,bacterial growth was reached at 50 h, however, maximum RuBisCOwas detected at 72 h after that it was declined. RuBisCO is alsoreported from Enteromorpha prolifera, Ulva lactuca, Chonrrus

Time (

0 20 40 60

Gro

wth

(OD

at 5

60 n

m)

0.0

0.1

0.2

0.3

0.4

0.5

GrowthRuBisCO activity

(a)

(b

Fig. 2. Growth and RuBisCO activity of Serratia sp. ISTDO4 isolated from bacterial commurepresents bacterial growth at O.D. at 595 nm with respect to time in hours (a) and y2 axmolecular weight markers of different molecular weight (14.4, 18.4, 25, 35, 45, 60.2 anbicarbonate and 2, crude protein lysate cross reacted RuBisCO (rbcL) antibody in Weste

crispus, Corallina officinalis, Dumontia incrassate and Porphyraumbilicalis 15.5, 9.4, 6.1, 52.7, 24.2 and 18.8 nmol CO2 fixed min�1 -mg�1 proteins respectively (Hilditch et al., 1991). RuBisCO activityof Serratia sp. ISTD04 was low as compared to those in higherplants showing the lower end value of 50 nmol CO2 fixed min�1 -mg�1 protein in Symphytum sp. and it was much lower than540–750 nmol CO2 fixed min�1 mg�1 protein in Nicotiana tabacumand 1700–2100 nmol CO2 fixed min�1 mg�1 protein in Spinaciaoleracea (Bahr et al., 1981).

The proteins of bacterial cell lysate were determined by SDS–PAGE after staining. The proteins present on gel surface transferredinto membrane filter and Western blot was performed. The pres-ence of RuBisCO protein was confirmed by SDS–PAGE and Westernblot analysis. Results of the study exhibited formation of clear bandof 55 kDa molecular weight size indicated cross reaction of proteinwith RuBisCO specific antibody recognized a polypeptide with amolecular weight of 55 kDa (Fig. 2b). Result confirmed the pres-ence of RuBisCO in Serratia sp. ISTD04. RuBisCO antibodies hadapproximate apparent molecular masses of 55 kDa. This proteinbands likely to be probable RuBisCO large-subunit polypeptidesdetected more prominently in the cell extract of Serratia sp.ISTD04.

h)

80 100 120 140

RuB

isC

O a

ctiv

ity n

anom

ole

CO

2 fix

ed m

g-1 m

in-1

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8

10

)

nity enriched in sodium bicarbonate (50 mM) in the chemostat. At right y1-axis, lineis activity of RuBisCO. SDS–PAGE for crude lysate of Serratia sp. In Lane, M is proteind 116 kDa); in lane 1, 5.6 lg crude lysate of induced bacteria with 50 mM sodiumrn blot analysis.

194 R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197

3.3. Analysis of hydrocarbons and lipids for production of biodiesel

The stationary phase culture was harvested, hydrocarbons wereextracted and the yield was 0.487 mg hydrocarbon mg�1 of cell dryweight. Hydrocarbons were identified by their M+ ions and com-parison of the mass spectra with those of the standards (Sigma)and also with the NIST library. The type of hydrocarbons producedby the Serratia sp. ISTD04 were analyzed by GC–MS and identifiedas saturated hydrocarbons in the range of C14–C24 and unsaturatedin the range of C13–C19 (Table 1). The saturated hydrocarbons pro-duced by the Serratia sp. ISTD04, pentadecane and docosane con-stituted 10.08% and 7%, respectively. 1-tridecene, 1-pentadecene,1-heptadecene and 1-octadecene were found as the major compo-nents among the unsaturated hydrocarbons produced by this bac-terium constituting 11.11%, 12.78%, 10.25% and 11.62%,respectively. Of the total hydrocarbons produced, total unsaturatedhydrocarbons in Serratia sp. ISTD04 constituted approximately51.86% and the total saturated hydrocarbon constituted approxi-mately 31.41% with the remaining amount 16% accounted to fattyalcohol, fatty aldehyde as well as other unidentified species.

Kerosene, jet fuel, and diesel fuel contain hydrocarbons in therange of C12–C20, whereas fuel oil has a range of hydrocarbons ofC20–C40. The hydrocarbons produced by Serratia sp. ISTD04 fallsin the range of C13–C24 which is equivalent to light oil that makeit a potential source of valuable fuel products like kerosene, jet fueland diesel fuel (Ladygina et al., 2006). Hydrocarbons falling underthis category have a clear advantage over gaseous fuel in their suit-ability for use with existing combustion and storage systems. Thestationary phase culture was harvested, lipids were extracted and

Table 1Composition of hydrocarbons of Serratia sp. ISTD04 isolated from bacterial community en

No. R.Time Compound name Molecular fo

1 14.135 1-Tridecene C13H26

2 14.294 Tetradecane C14H30

3 17.136 Pentadecane C15H32

4 18.823 1-Pentadecene C15H30

5 18.963 Hexadecane C16H34

6 20.238 Heptadecane C17H36

7 21.424 Octadecane C18H38

8 23.180 1-Heptadecene C17H34

9 23.294 Nonadecan C19H40

10 24.364 Docosane C22H46

11 25.835 Eicosane C20H42

12 27.160 1-Octadecene C18H36

13 27.249 Heneicosane C21H44

14 29.818 Tetracosane C24H50

15 30.791 1-Octadecanol C18H38O16 30.866 1-Tricosene C23H46

17 34.361 1-Nonadecene C19H38

Table 2Assignment of 1H NMR peak of lipid and FAME of Serratia sp. ISTD04 obtained from chemossource.

Proton(s) Functional group

CH3–C Terminal methyl group–(CH2)n– Backbone CH2

–CH2CH2COOH Beta methylene proton@CH–CH2– R-methylene group to one double bond–CH2COOH R-methylene group to acid–CH2COOR R-methylene group to ester@CH–CH2–CH@ R-methylene group to two double bonds–CH2OCOR Methylene group (C1 and C3) of glyceride–CHOCOR Methine proton at C2 of glyceride–CH@CH– Olefinic protons

the yield was 0.647 mg lipid mg�1 of cell dry weight culture. TheFTIR spectrum of bacterial lipids is shown in Fig. S1a. The promi-nent peaks of triacylglycerol due to C–H stretching mode in thewave number region of 2828 and 2929 cm�1, C@O stretching inthe region of 1743 cm�1 and C–O–C stretching and C–H bendingin the region of 900–1400 cm�1.

The 1H NMR spectrum of bacterial lipid is shown in Fig. S1b. Theproton resonances of the triacylglycerol of bacterial lipids are as-signed according to the literature data (Hopkins, 1966; Laetitiaet al., 2006). The olefinic protons –CH@CH– of unsaturated fattyacids resonate at 5.2–5.5 ppm, the multiplet is not baseline re-solved from the proton signal at 5.26 ppm (m) which was assignedto H-2 of the glycerol backbone (Table 2). The H-1 and H-3 protonsof glycerol resonate at 4.1 and 4.3 ppm, the assignments are inter-changeable. The H-2 and H-3 protons of acyl moieties in triacylgly-cerols resonate at 2.30 and 1.6 ppm, respectively, and the protonsof methylene envelope appear at 1.2 ppm. The methylene group(C1 and C3) of glyceride in triacylglycerols resonate at 4.03 and4.34 ppm, respectively revealing the presence of triacylglycerol inSerratia sp. ISTD04 (Laetitia et al., 2006).

The 13C NMR spectrum shows the resonances of carbons fromthe triglyceride fraction of bacterial lipid (Fig. S1c and d). The car-bon-13 resonances are grouped in four sets of signals, carbonyl car-bons resonating from 172 to 174 ppm, unsaturated carbonsresonate in the range from 124 to 134 ppm, glycerol backbone car-bons resonate from 60 to 72 ppm and aliphatic carbons resonatefrom 10 to 35 ppm. The 13C-NMR peaks at 173, 127.4, 69.2, 62.4,33.8, 31.9, 25.9 and 14.4 ppm of the carbons in triacylglycerol werereported (Laetitia et al., 2006).

riched in the chemostat in presence of sodium bicarbonate as sole carbon source.

rmula Relative molecular mass Relative content (%)

224 11.11198 1.08212 10.08210 12.78226 2.52240 2.38254 1.52238 10.25268 2.12310 6.97282 1.61252 11.62296 2.00338 1.00270 4.11322 0.98266 1.01

tat of bacterial community enriched in presence of sodium bicarbonate as sole carbon

Compound/chemical shift ppm

Fatty acid Lipids/TAG FAME

0.8–1.0 0.8–1.0 0.8–1.01.14–1.40 1.08–1.20 1.22–1.42

1.32–1.55 1.55–1.691.92–2.11 1.93–2.10 1.93–2.102.34 2.08–2.26

2.31 2.312.08–2.26 2.77 2.77– 4.09–4.34– 5.255.3–5.4 5.27–5.41 5.28–5.43

Table 3Fatty acid methyl esters in the biodieselformed after transesterification of lipids andhydrocarbons of Serratia sp. ISTD04 obtained from chemostat of bacterial communityenriched in presence of sodium bicarbonate as sole carbon source.

R. Time Area% Formula Name % FAME

12.372 4.40 C15H30O2 Tetradecanoic acid, methyl ester 5.5014.788 0.46 C17H34O2 Hexadecanoic acid, methyl ester 0.5715.227 1.20 C17H32O2 2-Hexadecenoic acid, methyl ester 1.5016.931 1.74 C16H32O3 Pentadecanoic acid, methyl ester 2.1717.310 35.73 C17H34O2 Methyl palmitate 40.6617.769 0.75 C17H32O2 9-Hexadecenoic acid, methyl ester 0.9320.218 26.86 C19H36O2 9-Octadecanoic acid, methyl ester 31.5721.782 0.92 C21H42O2 Eicosanoic acid, methyl ester 1.1521.975 8.91 C20H40O2 Nonadecanoic acid, methyl ester 5.0822.304 1.22 C19H36O2 9-Octadecanoic acid, methyl ester 1.5224.565 6.54 C20H38O2 10-Nonadecenoic acid, methyl ester 8.17

R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197 195

3.4. Optimization of transesterification reaction for production ofbiodiesel

Reaction conditions for transesterification, in terms of catalysttype, catalyst concentration, reaction temperature, reaction timeand rate of stirring, were optimized for maximum FAME yield.The most common basic catalysts KOH and NaOH were used in thisstudy. Experiments revealed that, the tested catalysts, sodiumhydroxide exhibited the highest yield of methyl esters during thetransestrerification of bacterial lipid with methanol. Theconcentrations of NaOH as catalyst, selected for this study were0.25–1.50% (on the basis of the weight of lipid) (Fig. 3a). In allthe reactions, conditions were maintained at 40 �C, catalyst con-centration of 1%, with an oil:methanol molar ratio of 1:6 at350 rpm for 3 h. To optimize the influence of the reaction, temper-ature on methyl ester formation, temperature variations adoptedin this study were 25, 30, 35, 40, 45, 50 and 55 �C (Fig. 3b).

The effect of stirring on FAME production was investigated infive experiments by differing stirring rates (150, 200, 250, 300and 350 rpm). The results clearly indicated that the optimumconcentration of NaOH required for effective transesterificationof bacterial lipid was 1.00% and 95% yield. After 3 h, the reactionwas completed up to 94%, 92%, and 88% at 40, 45, and 35 �C,respectively (Fig. 3c).

As depicted from Fig. 3d, a direct correlation was elucidatedbetween the stirring rate and FAME yield; i.e., as the rate of agita-tion was increased, an increase in yield was observed. Similarly, a

% of NaOH 0.2 0.4 0.6 0.8 1.0 1.2 1.4 1.6

Yiel

d of

FAM

E

0

20

40

60

80

100

120

Time1h 2h 3h 4h 5h 6h

Yiel

d of

FAM

E

0

20

40

60

80

100

120

(a)

(C)

Fig. 3. (a) Effect of the % NaOH concentration on yield of FAME, (b) effect of different tempof stirring (350 rpm rate of stirring, 1:6 lipid/methanol ratios, 3 h reaction duration and

mixing rate of 350 rpm afforded the optimum conversion of bacte-rial lipid to FAME (96%) after optimization of process parameters.

3.5. Determination of fatty ester profile and quality of biodiesel

The fatty acid methyl ester profile of biodiesel as determined byGC/MS is given in Table 3. The present GC–MS data also showedthe completion of the transesterification reaction. The FTIR spec-trum of FAME indicated most prominent peak is at 1733 cm�1

(C@O stretch), showing the presence of an ester. The band

Temperature25 30 35 40 45 50 55

Yiel

d of

FAM

E

0

20

40

60

80

100

120

RPM100 150 200 250 300 350

Yiel

d of

FAM

E

0

20

40

60

80

100

120

(b)

(b)

erature, (c) effect of time interval during transesterification and (d) effect of the rate45 �C reaction temperature was optimized for lipid production).

196 R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197

observed in the biodiesel at 1165 cm�1 is attributed to methylgroups near carbonyl groups (Roeges, 1994). The major changewas also observed at 2919 cm�1 which is characteristic of fatty acidmethyl esters as shown in Fig. S2(a). The 1H NMR spectrum ofFAME is shown in Table 2 and Fig. S2(b). The olefinic protons –CH@CH– of unsaturated fatty acids resonate at 5.2–5.5 ppm. Thea and b methylene group to ester in FAME resonate at 2.31 and1.55–1.69 ppm, respectively, and the protons of methyl group ofester appear at 3.67 ppm. The FTIR and NMR spectrum also reflectsthe conversion of triacylglycerols to methyl esters (Table 2).

The total unsaturated fatty acids found in Serratia sp. ISTD04and its methyl esters amount to approximately 55% and the totalsaturated fatty acids to approximately 45% as shown in Fig. S3.The characteristics of the FAME obtained from different optimizedmethods are presented in Table 3. The major FAMEs contained inthe biodiesel were esters of myristic acid (C14:0), palmitic acid(C16:0), oleic acid (C18:1), nonadecanoic acid, methyl ester (C19:0)and 10-nonadecenoic acid (C19:1). The most abundant compositionof bacterial lipids transesterified with methanol and base catalystis oleic acid methyl ester, which is suggested to according withthe standards of biodiesel. The sample consisted mainly of methylpalmitate and methyl oleate, which were almost 78% of the totalFAMEs. The gas chromatogram representing the fatty acid methylesters obtained from Serratia sp. ISTD04 indicated fatty acids suchas nonadecenoic acid and tetradecanoic acid constituted 8.17% and5.5%.

Oils containing higher level of saturated fatty acids than unsat-urated fatty acid may solidify and clog the fuel line during the win-ter condition (Demirbas, 2008). Biodiesel which contain high levelof unsaturated fatty acids are less viscous and show higher pourand cloud point properties which make biodiesel suitable for warmand cold weather conditions. It has been predicted that high levelsof oleic acid are best suited for biodiesel production (Robles-Med-ina et al., 2009). The use of biofuel in place of conventional fuelswould slow the progression of global warming by reducing emis-sion of carbon dioxide and hydrocarbon (Fjerbaek et al., 2009).FAME obtained from Serratia sp. ISTD04 contained higher propor-tion of oleic acid (33.57%) which makes it suitable for betterbiodiesel. Biodiesel compositions with polyunsaturated methyl es-ters are not suitable for vehicle use as they have low cetane num-bers and reduced oxidative stability, on the other hand saturatedfatty acids have higher cetane number (Knothe et al., 2011)(European standard EN 14214 limits to 12%). Thus, the FAMEs ob-tained from Serratia sp. ISTD04 make it suitable candidate to beused in the production of good quality biodiesel. Therefore, resultof the study indicated carbon dioxide concentrating chemolitho-trophic bacteria for production of biodiesel. Green algae have beenreported to sequester CO2, and evaluated for production of hydro-gen by Enterobacter cloacae IIT-BT 08 (Kumar et al., 2013). It is be-lieved that hydrogen is produced by enzyme hydrogenase but wehave no explanation at this stage whether Serratia sp. ISTD04 hascapability to produce hydrogen together with biodiesel as thestrain is not properly characterized.

4. Conclusion

CO2 induced production of carbonic anhydrase and RuBisCO re-vealed sequestration efficiency of Serratia sp. ISTD04 enriched inthe chemostat. The isolated strain can be a promising candidatefor research dedicated to exploitation of bacteria for lipid produc-tion to be used in production of biodiesel. Serratia sp. produced0.487 mg of hydrocarbons mg�1 cell dry weight and 0.647 mg of li-pid mg�1 dry cell weight. The hydrocarbons produced by Serratiasp. ISTD04 falls in the range of C13–C24 that make it a potentialsource of valuable fuel products like kerosene, jet fuel and diesel

fuel. The strain can be harnessed commercially to sequester CO2

into hydrocarbon and lipid fuel.

Acknowledgements

This research work was supported by the research grants ofDepartment of Biotechnology, Government of India, New Delhi, In-dia. The author (Srivastava S.) thanks University Grants Commis-sion (UGC), New Delhi, Government of India, for providing D.S.Kothari Post Doctoral Fellowship and Mr. Ajai Kumar of AdvancedInstrumentation Research Facility (AIRF) Jawaharlal Nehru Univer-sity, New Delhi for GC–MS, NMR and FTIR analysis.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at http://dx.doi.org/10.1016/j.biortech.2013.11.075.

References

Ashida, H., Danchin, A., Yokota, A., 2005. Was photosynthetic RuBisCO recruited byacquisitive evolution from RuBisCO-like proteins involved in sulfurmetabolism? Res. Microbiol. 156, 611–618.

Bahr, J.T., Johal, S., Bourque, D.P., 1981. High specific activity ribulose-1,5-bisphosphate carboxylase/oxygenase from Nicotiana tabacum. Photosyn. Res.2, 235–242.

De Muynck, W., Verbeken, K., De Belie, N., Verstraete, W., 2010. Influence of thecalcium dosage on the effectiveness of bacterially induced carbonateprecipitation on limestone. Ecol. Eng. 36, 99–111.

Demirbas, A., 2008. Biofuels sources, biofuel policy, biofuel economy and globalbiofuel projections. Energy Con. Manag. 49, 2106–2116.

Dou, Z., Heinhorst, S., Williams, E.B., Murin, C.D., Shively, J.M., Cannon, G.C., 2008.CO2 fixation kinetics of Halothiobacillus neapolitanus mutant carboxysomeslacking carbonic anhydrase suggest the shell acts as a diffusional barrier forCO2. J. Biol. Chem. 283, 10377–10384.

Fjerbaek, L., Christensen, K.V., Norddahl, B., 2009. A review of the current state ofbiodiesel production using enzymatic transesterification. Biotechnol. Bioeng.102, 1298–1315.

Hilditch, C.M., Jones, P.B., Balding, P., Smith, A.J., Rogers, L.J., 1991.Ribulosebisphosphate carboxylase/oxygenase from microalgae: proteolysisduring extraction and properties of enzyme from Porphyra umbilicalis.Phytochemistry 30, 745–750.

Hopkins, C.Y., 1966. Nuclear magnetic resonance in fatty acids and glycerides. Prog.Chem. Fats Other Lipids 8, 215–252.

Howarda, P., Middelhaufea, S., Moorea, K., Ednera, C., Kolaka, D.M., Taylora, G.N.,Parkera, D.A., Leea, R., Smirnoff, N., Avesa, S.J., Lovea, J., 2013. Synthesis ofcustomized petroleum-replica fuel molecules by targeted modification offree fatty acid pools in Escherichia coli. Proc. Natl. Acad. Sci. 110, 76376–77641.

Innocenti, A., Scozzafava, A., Parkkila, S., Puccetti, L., Simone, G.De., Supuran, C.T.,2008. Investigation of the esterase, phosphate, and sulfatase activities of thecytosolic mammalian carbonic anhydrase isoform I, II and XIII with 4-nitrophenyl esters as substrates. Bioorg. Med. Chem. Lett. 18, 2267–2271.

Jaiswal, P.K., Kohli, S., Gopal, M., Thakur, I.S., 2011. Isolation and characterization ofalkalotolerant Pseudomonas sp. strain ISTDF1 for degradation of dibenzofuran. J.Ind. Microbiol. Biotechnol. 38, 503–511.

Jo, B.H., Kim, I.G., Seo, J.H., Kang, D.G., Cha, H.J., 2013. Engineered Escherichia coliwith periplasmic carbonic anhydrase as a biocatalyst for CO2 sequestration.Appl. Environ. Microbiol.. http://dx.doi.org/10.1128/AEM.02400-13.

Knothe, G., Rashid, U., Yusup, S., Anwar, F., 2011. Fatty acids of Thespesia populnea:mass spectrometry of picolinyl esters of cyclopropene fatty acids. Eur. J. LipidSci. Technol. 113, 980–984.

Kumar, K., Roy, S., Das, D., 2013. Continuous mode of carbon dioxide sequestrationby C. sorokiniana and subsequent use of its biomass for hydrogen production byE. cloacae IIT-BT 08. Biores. Technol. 145, 116–122.

Kurian, D., Jansèn, T., Mäenpää, P., 2006. Proteomic analysis of heterotrophy inSynechocystis sp. PCC 6803. Proteome 6, 1483–1494.

Ladygina, N., Dedyukhina, E.G., Vainshtein, M.B., 2006. A review on microbialsynthesis of hydrocarbons. Proc. Biochem. 41, 1001–1014.

Laetitia, F., Fabien, R., Renaud, V., Fabiola, D., 2006. Chemical synthesis and NMRcharacterization of structured polyunsaturated triacylglycerols. Chem. Phys.Lipids 139, 125–136.

Lu, X., Vora, H., Khosla, C., 2008. Overproduction of free fatty acids in E. coli:implications for biodiesel production. Metab. Eng. 10, 333–339.

Mishra, M., Thakur, I.S., 2011. Purification, characterization and mass spectroscopicanalysis of thermo-alkalotolerant<beta>-1, 4 endoxylanase from Bacillus sp. andits potential for dye decolorization. Int. Biodeter. Biodeg. 65, 301–308.

R.K. Bharti et al. / Bioresource Technology 153 (2014) 189–197 197

Moazami, N., Ranjbar, R., Ashori, A., Tangestani, M., Nejad, A.S., 2011. Biomass andlipid productivities of marine microalgae isolated from the Persian Gulf and theQeshm Island. Biomass Bioenerg. 35, 1935–1939.

Murray, R., Badger, M.R., Price, G.D., 2003. CO2 concentrating mechanisms incyanobacteria: molecular components, their diversity and evolution. J. Exp. Bot.54, 609–622.

Park, M.O., Heguri, K., Hirata, K., Miyamoto, K., 2005. Production of alternatives tofuel oil from organic waste by the alkane-producing bacterium, Vibrio furnissiiM1. J. Appl. Microbiol. 98, 324–331.

Ramanan, R., Kannan, K., Sivanesan, S.D., Mudliar, S., Kaur, S., Tripathi, A.K.,Chakrabarti, T., 2009. Bio-sequestration of carbon dioxide using carbonicanhydrase enzyme purified from Citrobacterfreundii. World J. Microbiol.Biotechnol. 25, 981–987.

Robles-Medina, A., González-Moreno, P.A., Esteban-Cerdán, L., Molina-Grima, E.,2009. Biocatalysis: towards ever greener biodiesel production. Biotechnol. Adv.27, 398–408.

Rodriguez-Navarro, C., Rodriguez-Gallego, M., Ben Chekroun, K., González-Munoz,M.T., 2003. Conservation of ornamental stone by Myxococcus xanthus inducedcarbonate biomineralization. Appl. Environ. Microbiol. 69, 2182–2193.

Roeges, N.P.G., 1994. A Guide to Complete Interpretation of Infrared Spectra ofOrganic Structures. John Wiley and Sons, Chichester.

Rottig, A., Steinbüchel, A., 2013. Acyltransferases in bacteria. Microbiol. Mol. Biol.Rev. 77, 277–321.

Schmitt, J., Flemming, H.C., 1998. FTIR-spectroscopy in microbial and materialanalysis. Int. Biodeterior. Biodegrad. 41, l–11.

Schenk, P., Thomas-Hall, S., Stephens, E., Marx, U., Mussgnug, J., Posten, C., Kruse, O.,Hankamer, B., 2008. Second generation biofuels: high-efficiency microalgae forbiodiesel production. Bioenerg. Res. 1, 20–43.

Schirmer, A., Rude, M.A., Li, X., Popova, E., del Cardayre, S.B., 2010. Microbialbiosynthesis of alkanes. Science 329, 559–562.

Smith, K.S., Ferry, J.G., 2000. Procaryotic carbonic anhydrases. FEMS Microbiol. Rev.24, 335–366.

Tao, D., Wenkun, Z., 2012. Optimization of calcium carbonate precipitation forBacillus pausteurii. Appl. Mech. Mat. 178, 676–679.

Thakur, I.S., 1995. Structural and functional characterization of a stable, 4chlorosalicylic acid degrading, bacterial community in a chemostat. World J.Microbiol. Biotechnol. 11, 643–645.

Yeates, T.O., Kerfeld, C.A., Heinhorst, S., Cannon, G.C., Shivel, J.M., 2008. Protein-based organelles in bacteria: carboxysomes and related microcompartments.Nat. Rev. Microbiol. 6, 681–691.

Zamarreno, D.V., Inkpen, R., May, E., 2009. Influence of temperature onbiocalcification by non-sporing freshwater bacteria. Geomicrobiol. J. 26, 1–12.