International Biodeterioration & Biodegradation 53 (2004) 127–132www.elsevier.com/locate/ibiod

Extraction of carotenoid produced during methanolwaste biodegradation

Piotr Stepnowskia ;∗, Karl-Heinz Blotevogelb, Bernd Jastor/b

aFaculty of Chemistry, Waste Management Laboratory, University of Gda�nsk, ul. Sobieskiego 18, PL-80-952 Gdansk, PolandbCentre for Environmental Research and Technology UFT, University of Bremen, Leobener Str., Bremen D-28359, Germany

Received 29 July 2003; received in revised form 24 September 2003; accepted 16 October 2003

Abstract

In recent studies, we have developed an applied methanol recycling technology, designed for academic institutions utilizing chromato-graphic applications. The waste-free concept of the method includes biodegradation steps intended for further degradation of post-processingresidues. During the biodegradation process a natural pigment, a carotenoid, is biosynthesized in considerable amounts. In the present studya high-performance liquid chromatography–electrospray ionization–mass spectrometry (HPLC–ESI–MS) method was used to analyze andpreliminarily identify unknown pigments in the extracts of Methylobacterium organophylum used for methanol waste consumption. An-alytical results and theoretical considerations led us to propose carotenoid structures, namely dihydroxylycopene (dehydroxy derivativesof oscillol) or myxol, known to be isolated from bacteria.? 2003 Elsevier Ltd. All rights reserved.

Keywords: Biodegradation; Carotenoids; Methanol waste

1. Introduction

Recently, we have developed a recycling techniquefor chromatography solvents, or “total recycling”, bycombining a fractional distillation technique with thebioproduction of carotenoids as high value compounds(Stepnowski et al., 2002a, b). Our methodology developedfor methanol/water waste is based on a batch distillationsystem of the Hempel-type fractionating column. In thisprocedure, a certain amount of waste is produced at ?rst cutand in still bottoms that is not suitable for further recov-ery. This fraction is utilized in a biodegradation process.As methanol is ubiquitous in nature, it is readily degradedmostly by “methylotrophic” microorganisms, aerobicallyas well as anaerobically (Goldberg and Rokem, 1991). Ourresearch, therefore, focused on members of the so calledpink-pigmented, facultative methylotrophic bacteria, owingto their relatively easy culturing features, but most of all,their ability to assimilate methanolic redistillation wasteproducts yielding valuable carotenoids.

∗ Corresponding author. Tel.: +48-58-3450448; fax: +48-58-3410357.E-mail address: sox@chem.univ.gda.pl (P. Stepnowski).

Carotenoids are important natural pigments found widelyin microorganisms and plants. They are, of course, essen-tial to plants for photosynthesis, acting in light-harvestingand, especially, in protection against destructive photooxi-dation (Goodwin, 1984; Britton et al., 1995). Man requires asupply of �-carotene and related compounds for addition ascolourants to many manufactured foods, drinks and animalfeeds, either in the form of natural extracts or as pure com-pounds manufactured by chemical synthesis. Carotenoidsare, for example, used as natural food colourants or feed ad-ditives in aquaculture (Benemann, 1992; Torrissen, 1995).Several dietary studies have shown that carotenoids combatvarious types of cancer and other diseases because of theirantioxidant and/or provitamin A function (Astorg, 1997).The production of carotenoids by biotechnology is there-

fore of increasing interest, especially those from microbialsources such as the microalga Dunaliella salina or yeastslike Pha2a rhodozyma and Rhodotorula glutinis (Ramirezet al., 2000; Bhosale and Gadre 2001; Margalith, 1999). Inthis paper, a study on the identi?cation of a biosynthesizedcarotenoid produced during the biodegradation of methanolwaste is presented. Recently, several studies have been un-dertaken in carotenoid analysis involving high-performance

0964-8305/$ - see front matter ? 2003 Elsevier Ltd. All rights reserved.doi:10.1016/j.ibiod.2003.11.001

128 P. Stepnowski et al. / International Biodeterioration & Biodegradation 53 (2004) 127–132

Fig. 1. Chromatogram of saponi?ed extract of M. organophylium.

liquid chromatography (HPLC) coupled to mass detectorsusing continuous How-fast atom bombardment (CF-FAB),electrospray (ESI), and atmospheric pressure chemical ion-ization, or APCI (Van Breemen et al., 1996; Clark et al.,1996; Careri et al., 1999; Lacker et al., 1999). Therefore,we used HPLC–ESI–MS (Mass Spectroscopy), combiningit with diode-array technique, for structure elucidation.

2. Materials and methods

2.1. Biodegradation

Methylobacterium organophilum, strain DSMZ 760,was routinely grown in a standard medium containing5:0 g peptone and 3:0 g meat extract l−1 distilled wa-ter with 1% methanol (v/v) as carbon source at 28◦Con a rotary shaker. To maximize carotenoid produc-tion while biodegrading methanolic distillation residues(up to 3% v/v of methanol) a di/erent medium wasused, containing KNO3; 1:0 g; MgSO4:7H2O; 0:2 g.CaCl2:2H2O; 0:02 g; Na2HPO4; 0:23 g; NaH2PO4; 0:07 g;FeSO4:7H2O; 0:001 g; CuSO4:5H2O; 5:0 �g; H3BO3;10:0 �g, MnSO4:5H2O; 10:0 �g, ZnSO4:7H2O; 70:0 �g,MoO3; 10:0 �g; distilled water, 1 l. For the biotreatmentof the methanolic waste, the culture pH was maintained at6.8 under aerated conditions (4 l air min−1) and the tem-perature at 28◦C, and an impeller speed of 580 rpm wasused throughout. The growth rate was approx. 0:05 �m−1

in a fed-batch mode of 5-l bioreactor. The methanol wasconsumed within 3 days yielding a biomass of around 60 g(wet weight) which contained approx. 500 �g extracted

carotenoids g−1 cells. A detailed description is given inStepnowski et al. (2002a).

2.2. Extraction and saponi=cation

Since carotenoids are known to be esteri?ed by long chainfatty acids, a saponi?cation procedure was applied. Samplesof M. organophilum were centrifuged (× 5000 g), washedtwice with fresh culture medium and centrifuged again. Pel-lets were resuspended in methanol under a stream of nitro-gen in order to extract the carotenoids. Then samples werekept overnight at −20◦C under a nitrogen atmosphere. Themethanol extract was clari?ed by centrifugation and furtherprocessed. In order to hydrolyze carotenoid esters the ex-tracts were saponi?ed. To 5 ml extract, 2 ml 0:01 M NaOHin MeOH was added and the reaction mixture left for 8 h un-der a nitrogen atmosphere at room temperature in darkness.

2.3. LC–MS

For analysis of reaction mixtures by HPLC–ESI–MS,an HP 1100 series HPLC system (Hewlett-Packard)equipped with automatic sample injector, diode-array de-tector (DAD) and the HP 1100 series ESI mass selec-tivity detector was used. Separation was performed ona 250 mm × 4 mm × 10 �m LiChrosorb RP-18 column(LiChrosorb, Merck KGaA, Darmstadt, Germany) us-ing the isocratic solvent system developed by Yuan andChen (1998), acetonitrile:methanol (0:1 M ammoniumformate): dichloromethane (71:22:7, by volume). For elu-tion acetonitrile-G ChromasolvJ, Super Gradient Gradeand methanol-G ChromasolvJ (Riedel-de HaNen GmbH,

P. Stepnowski et al. / International Biodeterioration & Biodegradation 53 (2004) 127–132 129

Table 1Properties of analysed carotenoids. Comparison of retention times, absorbance maxima and molecular ions masses

Carotenoid Retention time (min) �max (nm) Molecular ion [M+H] Log P

Astaxanthin 4.34 480 597 13.26Extracted carotenoid (main peak) 4.65 462, 488, 520 569 —Lutein 5.02 429, 456, 485 569 14.82Canthaxantin 6.55 485 565 14.10�-carotene 10.15 427, 450, 475 537 17.62Spheroidene — — 569 16.921,1′- or 2,2′-dihydroxylycopene — — 569 14.712′-dehydroxymyxol — — 569 14.83

Fig. 2. Spectrum of the carotenoid extracted from M. organophylium.

Fig. 3. Molecular peak ion of the carotenoid extracted from M. organophylium.

130 P. Stepnowski et al. / International Biodeterioration & Biodegradation 53 (2004) 127–132

OH

OH

OH

OH

OH

OH

O

OH

OH

O

OMe

OH

OH

O

O

OH

OH

2,2'dihydroxylycopene (m/z 568)

2'dehydroxy myxol (m/z 568)

Astaxanthin m/z 596 (3R, 3'R)

spheroidene (m/z 568)

Zeaxanthin (m/z 568)

Lutein (m/z 568)

β,β− carotene (m/z536)

canthaxantin (m/z 564)

1,1'dihydroxylycopene (m/z 568)

Fig. 4. Structures of analysed and hypothetical carotenoids (semi-systematic names).

Germany) dichloromethane (Fluka Chemika AG Buchs,Switzerland) and ammonium formate (Merck KGaA,Darmstadt, Germany) were used. The How rate was set to0:7 ml min−1, and injection volume was 10 �l. The DADwas operated at 450 and 480 nm simultaneously. Operatingparameters of the ESI–MS were optimized using the direct

infusion of a astaxanthin standard solution in the mobilephase: positive ion mode, dry gas How (N2)10 l min

−1,nebulizer pressure 50 psi, drying gas temperature 350◦Cand capillary voltage at 3500 V. Mass spectra were ac-quired over the scan range m/z 500–600 using a 0.1 unitstep size.

P. Stepnowski et al. / International Biodeterioration & Biodegradation 53 (2004) 127–132 131

Carotenoid standards were separated within one chro-matographic run, thus obtaining retention times and visi-ble absorption spectra. Molecular ion peaks were detectedfor �-carotene (m/z 537 [M+H]), canthaxanthin (m/z 565[M+H]), lutein (m/z 569 [M+H]) and astaxanthin (m/z 597[M+H]) (Fig. 4).

2.4. Determination of hydrophobicity

Partition coeRcient values (log P) of all studied com-pounds were calculated from the structures using fragmentconstant estimation methodology (KowwinJ software ver-sion 1.66) developed at Syracuse Research Corporation(Meylan and Howard, 1995).

3. Results and discussion

Direct analysis of the extract showed a complex pattern ofunidenti?able carotenoids. None of the known carotenoidsin the free form could be detected. Applying the saponi?ca-tion procedure led to an e/ective hydrolysis of all derivativesyielding two peaks (Fig. 1). The retention times were foundto be between those of astaxanthin and lutein standards (re-tention times, absorbance maxima and molecular ion peaksmasses are given in Table 1). The photodiode array analy-sis showed a ?ne structured 3 maxima shape type spectrum(Fig. 2). This excludes the presence of cyclic ketones suchas astaxanthin or canthaxanthin, which are known to haveonly one maximum. For the minor HPLC peak a 380 nm ab-sorption band is registered additionally which is character-istic for cis-isomers in the family of carotenoids. The ESI–MS analysis exhibited a molecular ion at m/z 569[M+H]for both peaks (Fig. 3). This mass corresponds to the stan-dards of lutein (or zeaxanthin). Comparing the visible spec-tra, lutein and the analysed compound show a similar pat-tern, but the bacterial sample has a signi?cant bathochromicshift. This is con?rmed by the red colour of the studied ex-tract, while lutein (or zeaxanthin) solutions are yellow. Asolvent e/ect can also be excluded, since all spectra weretaken with a photodiode array using the same mobile phase.Owing to the lack of standards for rare carotenoids, fur-ther consideration was made on a theoretical basis. Redcolouration is due to absorption in the higher visual range,and the III maximum of the extracted carotenoid is wellabove 500 nm. This is most probably due to the presenceof more conjugated trans-oriented double bonds because ofthe acyclic character of the unknown carotenoid. We thenchecked whether this hypothesis was in accordance withthe observed lipophilicity of the bacterial product. Acyclicand non-substituted compounds such as bacterial lycopenewould have a much longer time of interaction with RP-18phase used. Acyclic diols with more �-electrons are there-fore in accordance with our assignment of structure. Amongtypical microbial carotenoids given by Britton et al. (1995)only a few of the C40 type have a III maximum greater than

500 nm. Fig. 4 shows four possible structures of them/z 569[M+H]. Owing to a calculated hydrophobicity, the retentionof methoxy compound (spheroidene) is improbable. Mostlikely structures are the 1,1′- and 2,2′-didehydroxy oscillolas well as 2′-dehydroxymyxol acyclic only on one terminus.These two structures are in agreement with a chromophorewith enough conjugated double bonds to yield observed vis-ible spectrum and are hydrophilic enough to be eluted closeto a congeneric dihydroxy group-containing lutein, zeaxan-thin and astaxanthin. Because of the molecular ion identi-?ed we can exclude C30, C45 and C50 compounds, becauseonly C40 pigment ?ts this spectrum. Accordingly, we pro-pose the molecular formula of C40H56O2 and suggest themost presumptive three-dimensional structures as 1,1′- or2,2′-dihydroxylycopene (didehydroxy derivative of oscillol)or dehydroxy derivative of myxol.In conclusion, our assigned structure leads us to propose

that M. organophilum synthesizes, as a main product, anacyclic all-trans carotenoid. The minor peak could eitherbe a cis form of such a compound synthesized by bacteriaor formed during the extraction and saponi?cation proce-dure. To verify the proposed structure, further studies byhigh-resolution mass spectrometry and 1H−NMR analysiswill be required.

Acknowledgements

This study was partially supported by the Polish Ministryof Scienti?c Research and Information Technology underthe Grant DS 8390-4-0141-3. Assistance of Dr J. Ranke inpreparation of this manuscript is greatly acknowledged.

References

Astorg, P., 1997. Food carotenoids and cancer prevention: an overview ofcurrent research. Trends in Food Science and Technology 8, 406–413.

Benemann, J.R., 1992. Microalgae aquaculture feeds. Journal of AppliedPhycology 4, 233–245.

Bhosale, P., Gadre, R.V., 2001. T-carotene production in sugarcanemolasses by a Rhodotorula glutinis mutant. Journal of IndustrialMicrobiology and Biotechnology 26, 327–332.

Britton, G., Liaaen-Jensen, S., Pfander, H., 1995. Carotenoids— Isolationand Analysis. BirkhNauser, Basle.

Careri, M., Elviri, L., Mangia, A., 1999. Liquid chromatography–electronspray mass spectrometry of �-carotene and xanthophylls.Journal of Chromatography A 854, 233–244.

Clark, P.A., Barnes, K.A., Startin, J.R., Ibe, F.I., Shepherd, M.J.,1996. High performance liquid chromatography/atmospheric pressurechemical ionisation mass spectrometry for the determination ofcarotenoids. Rapid Communications in Mass Spectrometry 10, 1781–1785.

Goldberg, I., Rokem, J., 1991. Biology of Methylotrophs. Butterworth–Heinemann, Stoneham, MA.

Goodwin, T.W., 1984. The Biochemistry of Carotenoids. Chapman andHall, New York.

Lacker, T., Strohschein, S., Albert, K., 1999. Separation and identi?cationof various carotenoids by C30 reverse phase HPLC coupled to UV andatmospheric pressure chemical ionisation mass spectrometric detection.Journal of Chromatography A 854, 37–44.

132 P. Stepnowski et al. / International Biodeterioration & Biodegradation 53 (2004) 127–132

Margalith, P.Z., 1999. Production of ketocarotenoids by microalgae.Applied Microbiology and Biotechnology 51, 431–438.

Meylan, W.M., Howard, P.H., 1995. Atom/fragment contributionmethod for estimating octanol–water partition coeRcients. JournalPharmaceutical Science 84, 83–92.

Ramirez, J., NuVnez, M.L., Valdivia, R., 2000. Increased astaxanthinproduction by a Pha2a rhodozyma mutant grown on date juice fromYucca =llifera. Journal of Industrial Microbiology and Biotechnology24, 187–190.

Stepnowski, P., Blotevogel, K.-H., Jastor/, B., 2002a. Applied waste-freerecovery of methanol. A sustainable solution for chromatographylaboratories. Environmental Science and Pollution Research 9,34–38.

Stepnowski, P., Blotevogel, K.-H., Ganczarek, P., Fischer, U.,Jastor/, B., 2002b. Total recycling of chromatographic solvents—applied management of methanol and acetonitrile waste. ResourceConservation and Recycling 35, 163–175.

Torrissen, O.J., 1995. Strategies for salmonid pigmentation. Journal ofApplied Ichthyology 11, 276–281.

Van Breemen, R.B., Huang, C.R., Tan, Y., Sander, L.C., Schilling,A.B., 1996. Liquid chromatography/mass spectrometry of carotenoidsusing atmospheric pressure chemical ionisation. Journal of MassSpectrometry 31, 975–981.

Yuan, J.-P., Chen, F., 1998. Chromatographic separation and puri?cationof trans-astaxanthin from extracts of Haematococcus pluvialis. Journalof Agricultural and Food Chemistry 46, 3371–3375.