Microalgae mediated photoproduction of b-carotene in aqueous�/

organic two phase systems

Rosa Leon a,*, Marta Martın a, Javier Vigara a, Carlos Vilchez a, Jose Marıa Vega b

a Dpt. de Quımica y CC MM (Area de Bioquımica), Facultad de Ciencias Experimentales, Campus del Carmen, Universidad de Huelva, 21071 Huelva,

Spainb Dpt. Bioquımica Vegetal y Fotosıntesis, Facultad de Quımica, Universidad de Sevilla, Apdo. 553, 41012 Sevilla, Spain

Biomolecular Engineering 20 (2003) 177�/182

www.elsevier.com/locate/geneanabioeng

Abstract

Improving productivity is a usual requirement for most biotechnological processes, and the utilisation of two-phase aqueous

organic systems has proved to be an effective way to improve the productivity of poorly water-soluble or toxic compounds. The high

hydrophobicity of b-carotene, which is highly demanded by the pharma and agrofood industry, makes it a good candidate for

aqueous/organic biphasic photoproduction. In the present work we have investigated the viability of a two-phase system for the

production of b-carotene by the marine microalgae Dunaliella salina using decane as organic phase. Decane, with a log Poctanol value

of 5.6, showed no toxicity to Dunaliella cells for more than 72 h, and its ability for b-carotene extraction is acceptable. Transferring

Dunaliella cells from standard to carotenogenic conditions caused inhibition of chlorophyll production and induced a strong

synthesis of b-carotene. The two-phase aqueous/decane system was stable and b-carotene content of the cells was increasing during

4-days. About 8% of the total carotenoids produced were excreted and extracted into the decane phase.

# 2003 Published by Elsevier Science B.V.

Keywords: b-Carotene; Dunaliella salina ; Biphasic systems; Decane

1. Introduction

In the last years an increasing interest for microalgae

biotechnology has emerged due to the variety of

biotechnologically interesting compounds that micro-

algae can synthesise. Food additives, colorants, fluor-

escent compounds, antioxidants, isotopically labelled

compounds are only some examples of the potentiality

of microalgae for the production of bioactive natural

compounds [1,2]. Despite the ability of microalgae to

produce many compounds of interest and their potential

as a source of new unexplored natural products, the

biotechnological applications of microalgae have been

less studied than those involving other microorganisms.

Improving productivity is a usual requirement for

most biotechnological processes, and the use of two-

phase aqueous organic systems has proved to be an

effective way to improve the productivity of poorly

water-soluble or toxic compounds. In these systems a

biocompatible organic solvent is in contact with the

aqueous phase where the cells are carrying out the

bioconversion. The product is continuously extracted

into the organic phase overcoming the low solubility of

the product, avoiding the possible toxic effect of the

product and facilitating product recovery and contin-

uous operation [3]. Biphasic aqueous/organic systems

have been widely applied in bioconversions catalysed by

bacteria [4,5], but only few examples of two-phase

applied systems for plant cells [6,7] or microalgae [8]

have been described.

Carotenoids are a wide family of isoprenoids with 40

carbon atoms comprising carotenes and their oxyge-

nated derivates, the xanthophylls. They act as secondary

pigments in photosynthetic organisms, but also play an

important role as antioxidants and provitamin factors

for non-photosynthetic organisms that must include

them in their diet. Their antioxidant and colorant

properties, which are related to the system of conjugated

double bonds, are responsible for the therapeutic,

dietetic and industrial applications of carotenoids.

* Corresponding author. Tel.: �/34-959-01-9951; fax: �/34-959-01-

9942.

E-mail address: rleon@uhu.es (R. Leon).

1389-0344/03/$ - see front matter # 2003 Published by Elsevier Science B.V.

doi:10.1016/S1389-0344(03)00048-0

b-Carotene is highly demanded by the pharma and

agrofood industry. It is commercialised as dietetic

compound, food additive (colorant, antioxidant), ani-

mal feed, etc. The chlorophyte microalgae Dunaliella

salina is the main source of natural b-carotene and

several industrial exploitations of Dunaliella are opera-

tional in Australia, Israel and USA for this purpose [9].

Nevertheless, and in spite of the multiple natural sources

of b-carotene, more than 90% of commercialised b-

carotene is chemically synthesised. New approaches to

improve productivity of biosynthetic processes and

make them competitive against chemical synthesis arenecessary.

The high hydrophobicity of b-carotene makes it a

good candidate for aqueous/organic biphasic photopro-

duction. Based on previous studies on organic solvent

toxicity on several microalgae strains [10] we have

investigated the viability of a two-phase system for the

production of b-carotene using decane as organic phase.

2. Materials and methods

2.1. Microorganism and culture conditions

D. salina (UTEX 2538) was kindly provided by the

Plant Biochemistry and Photosynthesis Institute (CSIC,

Seville). Standard cultures were grown in mineral liquid

medium at 25 8C, bubbled with air containing 5% (v/v)

CO2 and continuously illuminated with cool white and

daylight from fluorescent lamps (100 mE m�2 s�1). The

composition of the liquid medium was the described byJohnson et al. [11]. For carotenogenesis induction D.

salina cells were cultured in the same liquid medium

without nitrogen source.

2.2. Microalgae culture in aqueous/organic biphasic

systems

Aqueous/organic biphasic cultures were carried out in

the same conditions described above except that an

organic phase of decane was added. The phase ratio was

2:1 (aqueous�/organic). The percentage of organic

solvent was ensured by periodical addition of the solvent

to compensate its loss by evaporation when necessary.

2.3. Cell viability

Light dependent photosynthetic activity was used to

quantify the cell viability. For photosynthetic activity

determinations 1 ml microalgae culture was placed in

the reaction chamber of a Clark-type electrode and thelight-dependent O2-evolution was measured. Measure-

ments were made at 25 8C under saturating white light

(1500 mE m�2 s�1).

2.4. Analytical determinations

Total chlorophyll, chlorophyll a, chlorophyll b and

total carotenoids were extracted with 80% acetone anddetermined by spectrophotometric methods using the

equations proposed by Lichtenthaler [12] or by HPLC

analysis (see below). Protein content was determined

with the Bio-Rad protein assay dye reagent following

the instructions of the manufacturer (Bio-Rad, USA)

and using bovine serum albumin as standard. All

experimental data are the average of at least two

replicates (standard deviation, sB/3%).

2.5. Specific growth rate determination

The logarithm of the number of cells, measured by

counting in a Neubauer chamber, was plotted against

the time over 72 h and the specific growth rate wascalculated from the slope of this plot.

2.6. HPLC analysis of pigments

The separation and chromatographic analysis of

pigments was preformed in a Merck Hitachi HPLCequipped with a UV�/Vis detector as described by

Young et al. [13], using a RP-18 column and a flow

rate of 1 ml min�1. The mobile phase consisted on:

solvent A, ethyl acetate; solvent B acetonitrile/water

(9:1, v/v) and the gradient programme applied was: 0�/16

min, 0�/60% A; 16�/30 min, 60% A; 30�/35 min, 100%.

Pigments detection was carried out at 450 nm, and their

identification and quantification was achieved by inject-ing known amounts of pigments standards (chlorophyll

a, chlorophyll b, lutein and b-carotene) supplied by

SIGMA.

2.7. Solvent biocompatibility assays

In solvent toxicity experiments microalgae cultures

were incubated for 15 min in 50 ml of growth medium

supplemented with 5% (v/v) of the indicated organic

solvent and vigorously stirred for several minutes to

ensure saturation of the aqueous medium with the

solvent.

2.8. b-Carotene solubility assays

Saturated solutions of b-Carotene in the different

organic solvents were obtained adding amounts of b-

carotene above the solubility limit to 5 ml of each

organic solvent, stirring vigorously with a vortex during

several minutes followed by 200 rpm shaking during 2 hat 25 8C. After centrifugation and filtration through a

syringe filter with 0.22 mm of pore size, b-carotene

content in saturated solutions was determined as

R. Leon et al. / Biomolecular Engineering 20 (2003) 177�/182178

indicated above, using molar absorption coefficients

calculated for b-carotene in each organic solvent.

2.9. Pigments extraction ability assays

The ability of different solvents to extract pigments

from inside D. salina cells was estimated incubating 5-ml

Dunaliella cultures with 2 ml of the indicated organic

solvent and shaking at 200 rpm during 24 h at 25 8C.

After this time, tubes were centrifuged and total

carotenoid content in the organic phase was determinedas indicated above.

3. Results and discussion

3.1. Choice of biocompatible organic solvent

The toxic effect caused by some organic solvents on

living cells can be attributed to the contribution of

molecular and phase toxicity. Solvent molecular toxicity

is caused by the solvent molecules dissolved in the

aqueous phase, and the phase toxicity is due to direct

contact of the organic phase with the cells and becomes

more important at high agitation speeds [14,15]. Toevaluate the effect of toxicity caused by solvents of the

alkanols and alkanes series on the microalgae D. salina

biocompatibility assays were carried out as indicated in

Section 2 avoiding emulsification of liquid phases. In

these conditions molecular toxicity is the main contri-

butor to the toxicity of the solvent. The relative activity

of the cells was correlated with the log Poctanol, an

indicator of the hydrophobicity of these solvents.Decane, with a log Poctanol equal to 5.6, was the most

polar solvent that D. salina cells were able to tolerate

without an appreciable degradation of its growth rate or

photosynthetic activity for 72 h (Table 1). For isooctane

and all solvents with log Poctanol lower than 5.6 there was

an important loss of viability.

The pattern of solvent toxicity against hydrophobicityof the solvent observed in Dunaliella is exactly the same

sigmodial curve observed for bacteria [16], plant [7] and

other microalgae [10]. Solvents with a log Poctanol value

higher than the inflection point of this curve are so

hydrophobic that their solubility in water is not enough

to reach the minimum aqueous concentration that

causes toxicity. On the contrary, solvents with log Pocta-

nol value under the inflection point reach the toxicconcentration easily and are not biocompatible. The

value of this limiting log Poctanol value can be very

different for different cells. In a previous work we tested

the toxic effect of several organic solvents on some

representative photosynthetic microorganisms [10]. And

we observed that the tolerance of microalgae was

intermediate between that reported for bacteria and

for plant cell suspensions. Solvent tolerance observedfor Dunaliella , with a minimum log Poctanol value

tolerated of 5.6 is very similar to that reported for other

microalgae and lower than that reported for the

cyanobacteria Anabaena [10].

Comparison with data reported by other authors is

not easy because of the different biocompatibility

criteria adopted. Most authors choose the metabolic

activity directly involved in the bioconversion of interest[7,17] or growth parameters [18,19]. Hejazi et al. [20]

also studied the toxicity of alkanes solvents in Dunaliella

cells using the photosynthetic activity as metabolic

activity to define biocompatibility of the solvents.

They found a similar general response, although the

limiting log Poctanol value was higher than that we

previously reported [10]. They do not give any explana-

tion for this apparent disagreement, but we thinkdifferent culture conditions, such as the agitation

mode, can greatly influence on the viability of the cells

and explain it. All our experiments were done on small

bubbled flasks, while they used stirred bottles. Intense

mixing of aqueous and organic will enhance extraction

but can enhance damage caused by organic solvent, so a

compromise between toxicity and extraction ability has

to be taken into account when choosing the organicphase.

3.2. Effect of the presence of an organic phase on

carotenoids production by Dunaliella salina

It is generally accepted that D. salina synthesises big

quantities of b-carotene when subjected to stressing

conditions such as nutrient starvation, high light in-

tensity, extreme temperatures or salinity. Nitrogenstarvation induced a 33-fold increase of the carotene/

chlorophyll ratio, from 0.09 to approximately 3, which

is evidenced by the orange colour acquired by Dunaliella

Table 1

Toxicity of organic solvents of the alkanes serie on D. salina

log Poctanol m (h�1) Photosynthetic activity (%)

Control �/ 0.012 100

Pentane 3 0 0

Hexane 3.5 0 0

Heptane 4 0 0

Isooctane 4.5 0 29

Decane 5.6 0.011 111

Undecane 6.1 0.012 95

Dodecane 6.6 0.012 110

Tetradecane 7.6 0.013 97

Hexadecane 8.8 0.012 103

50-ml of a D. salina cell suspension centrifuged and resuspended in

solvent saturated growth medium supplemented with 5% (v/v) of the

indicated organic solvent. The cells were cultured in this medium and

samples were periodically withdrawn for photosynthetic activity and

number of cells determination. Specific growth over a period of 72 h

and mean photosynthetic activity value over the same period are

shown.

R. Leon et al. / Biomolecular Engineering 20 (2003) 177�/182 179

cells and from the relative height of peaks in the HPLC

chromatograms (Fig. 1). The presence of the organic

phase itself can be a stressing factor that enhances

carotenoids synthesis. To investigate this possibility cells

of D. salina were cultured in the presence of decane,

which log Poctanol value is around the limiting toxicity

value for D. salina cells as shown before (Table 1).Cells cultured in standard conditions were resupended

in fresh medium and cultured for 72 h in the presence of

a second phase of the decane 5% (v/v). Chlorophyll and

b-carotene content and photosynthetic activity of the

cells were followed and compared with control cells

grown under standard conditions.

In Fig. 2 can be seen that Dunaliella cells grown in the

presence of decane showed almost the same chlorophyll

content and slightly lower photosynthetic activity value

than control cells. Only a slight induction of carotene

synthesis in the presence of decane is observed, about

20% higher content of b carotene for cells grown in the

presence of decane, which is far from the 33-foldincrease observed when Dunaliella cells are subjected

to nitrogen starvation. Nevertheless we can confirm that

decane, is not toxic to viability of the cells and, also does

not affect the synthesis of carotene, which is enhanced.

Similar induction was reported by Hejazi et al. [20] for

Dunaliella cells in the presence of dodecane.

3.3. Extraction ability of different organic solvent

b-Carotene is synthesised by most plant tissues and is

mainly located in the thylakoid membrane of the

chloroplast where it acts as an accessory photosynthetic

pigment. In D. salina where the concentration of b-carotene under stressed conditions can reach really high

Fig. 1. Separation of pigments of control (A) and stressed (B) D. salina cells by HPLC. Stressed cells were subjected to nitrogen starvation during 96

h. The pigments composition of decane phase after 96 h in contact with a culture of stressed D. salina cells is also shown (C). Conditions for

chromatographic separation are described in Section 2. Peaks were identified as: lutein (1), chlorophyll b (2), chlorophyll a (3) and b-carotene (several

isomers) (4).

Fig. 2. Effect of decane on D. salina cells. Chlorophyll (A) and b-carotene (B) content and photosynthetic activity (C) of cells grown in control

conditions ( ) and in the presence of decane 5% (v/v) (I) were followed for 72 h.

R. Leon et al. / Biomolecular Engineering 20 (2003) 177�/182180

values it has been shown that b-carotene accumulates

forming liphophylic globules that play a protective role,

acting against free-radical and photoinhibitory pro-

cesses [21]. Excretion of b-carotene into the culture

medium is very low under normal conditions. Therefore,

in order to establish an aqueous organic two-phase

system with viable cells, the solvent chosen as organic

phase must satisfy certain minimum requirements.

Sufficient solubility for b-carotene in the chosen solvent,

the ability to pass through the plasma membrane,

extracting b-carotene and transporting it into the culture

medium without toxic effects and, if possible, sufficient

selectivity.

To have a global view of the properties of different

alkanols and alkanes we calculated the solubility of

commercial b-carotene in these solvents and determined

their ability to extract b-carotene from carotenogenesis-

induced Dunaliella cells as described in Section 2. The

results are shown in Fig. 3 for several solvents ordered

by increasing log Poctanol value. Data about the biocom-

patibility of these solvents, expressed as the percentage

of photosynthetic activity kept after 15 min of incuba-

tion with the solvent, are also shown.

The extractive ability and the biocompatibility of thesolvents studied show opposite trends (Fig. 3). More

hydrophobic solvents*/higher log Poctanol*/are less

toxic for the microalgae but carotenoids extraction

from the cells is less effective. However, the solubility

of b-carotene, which is a very hydrophobic compound

(log Poctanol�/17), is higher in the alkanes than in the

alkanols. We can consequently affirm that the plasmatic

membrane of Dunaliella cells is the main barrier for theextraction of carotenoids by the less toxic alkanes.

Decane with a log Poctanol value of 5.6 is not toxic to

Dunaliella cells. b-Carotene is very soluble in decane, as

in all other alkanes, and its ability for b-carotene

extraction from cells, although much lower than that

of the toxic alkanols is acceptable, about 20% of the

maximum and 2-fold the extraction ability of other

alkanes with higher log Poctanol value. Decane, as allother alkanes, is able to extract more efficiently the

hydrocarbonated b-carotene than the chlorophylls or

oxygenated xantophylls guaranteeing the selectivity of

the extraction, as shown in Fig. 1. Similar selectivity was

reported when using other alkanes, such as dodecane

[20].

3.4. Discontinuous production of b-carotene by D. salina

cells in the presence of the biocompatible solvent decane

Decane is not toxic for D. salina cells and is a good

and selective solvent for b-carotene, so it is a good

candidate to be used as organic phase in an aqueous/

organic biphasic system for the production of carotenes

by microalgae. Cells of D. salina grown in standardconditions were harvested by centrifugation, resus-

pended in fresh nitrogen free medium and cultured in

a biphasic system with decane as organic phase. The

phase ratio was 2:1 (aqueous�/organic). Carotene and

chlorophyll evolution in this system was followed for 4

days (Fig. 4c) compared with a culture in carotenogenic

Fig. 3. Extraction ability, solubility and biocompatibility of different

solvents on D. salina cells. Extractive ability (j), biocompatibility (I)

of different solvents on D. salina cells and solubility ( ) of b-carotene

in these solvents, measured as explained in Section 2, are shown.

Solvents are ordered by increasing log Poctanol value. 100% is the

maximum value observed for solubility, extraction and photosynthetic

activity, respectively.

Fig. 4. Total carotenoids (j) and chlorophyll (") content in D. salina cells cultured under normal conditions (A), under normal conditions without

nitrogen (B) and in a biphasic system without nitrogen (C) was followed during several days. The biphasic system consisted on 200 ml of (N-)culture

medium and 100 ml of decane. The evolution of carotenes in the organic phase (') of the biphasic system is also shown.

R. Leon et al. / Biomolecular Engineering 20 (2003) 177�/182 181

conditions without organic phase (Fig. 4b) and with a

control culture in standard conditions (Fig. 4a).

Transferring Dunaliella cells from standard to caro-

tenogenic conditions caused inhibition of chlorophyllproduction and induced a strong synthesis of b-caro-

tene. The carotenes/chlorophyll ratio increased 30-fold

in 4 days in nitrogen free cultures, both with and

without decane. Slight carotenegenesis induction that

we observed for cells cultured with decane (Fig. 2) is

hidden by the big induction due to the deficiency of

nitrogen source. The system was stable and b-carotene

content of the cells was increasing during 4-days. In thebiphasic system about 8% of the total carotenoids

produced were excreted and extracted into the decane

phase. For 4 days the cells were viable and produced b-

carotene. Furthermore the extraction is very selective,

especially in relation to chlorophylls and xantophylls

(see Fig. 1). The use of additional permeabilisation

agents to increase extraction is now under investigation.

Acknowledgements

R. Leon gratefully acknowledges a contract from the

Spanish Ministry of Science and Technology within the

‘‘Ramon y Cajal’’ Research Programme.

References

[1] Yamaguchi K. J Appl Phycol 1997;8:487�/502.

[2] Vılchez C, Garbayo I, Lobato MJ, Vega JM. Enzyme Microbiol

Technol 1997;20:562�/72.

[3] Laane C, Boeren S, Vos K, Veeger C. Biotechnol Bioeng

1987;30:81�/7.

[4] Leon R, Fernandes P, Pinheiro HM, Cabral JMS. Enzyme

Microb Technol 1998;23:483�/500.

[5] Fernandes P, Cabral JMS, Pinheiro HM. Enzyme Microb

Technol 1995;17(2):163�/7.

[6] Dutta A, Pedersen H, Chin CK. Ann New York Acad Sci

1994;745:251�/60.

[7] Bassetti L, Pijnenburg J, Tramper J. Biotechnol Lett

1996;18:377�/82.

[8] Largeau C, Bailliez C, Yang LW, Frenz J, Casadevall E. In:

Stadler T, Mollion, Verdus, Karamanos, Morvan, Christiaen,

editors. Algal biotechnology. London: Elsevier, 1988:245�/53.

[9] Borowitzka MA. J Biotechnol 1999;70:313�/21.

[10] Leon R, Garbayo I, Hernandez R, Vigara J, Vılchez C. Enzyme

Microbiol Technol 2001;29:173�/80.

[11] Johnson MK, Johnson EJ, McElroy RD, Speer HL, Bruff BS. J

Bacteriol 1968;95:1461�/8.

[12] Lichtenhaler HK. Methods Enzymol 1987;148:350�/82.

[13] Young A, Orset S, Tsavalos A. In: Pessarakli M, editor. Hand-

book of photosynthesis. New York: Marcel Dekker, 1997:597�/

622.

[14] Sikkema J, Bont JAM, Poolman B. Microbiol Rev 1995;59:201�/

22.

[15] Bar R. J Chem Technol Biotechnol 1998;43:49�/62.

[16] Vermue M, Sikkema J, Verheul A, Bakker R, Tramper J.

Biotechnol Bioeng 1993;42:747�/58.

[17] Osborne SJ, Leaver J, Turner MK, Dunnill P. Enzyme Microbiol

Technol 1990;12:281�/91.

[18] Rajagopal AN. Enzyme Microbiol Technol 1996;19:606�/13.

[19] Inoue A, Horikoshi K. J Ferm Bioeng 1991;3:194�/6.

[20] Hejazi MA, Lamarliere C, Rocha JMS, Vermue M, Tramper J,

Wijffels RH. Biotechnol Bioeng 2002;79(1):30�/6.

[21] Ben-Amotz A, Katz A, Avron M. J Phycol 1982;18:529�/37.

R. Leon et al. / Biomolecular Engineering 20 (2003) 177�/182182