Harvesting of microalgae by bio-flocculation

Abstract

The high-energy input for harvesting biomass makes current commercial microalgal biodiesel production economically unfeasible. A novel harvesting method is presented as a cost and energy efficient alternative: the bio-flocculation by using one flocculating microalga to concentrate the non-flocculating microalga of interest. Three flocculating microalgae, tested for harvesting of microalgae from different habitats, improved the sedimentation rate of the accompanying microalga and increased the recovery of biomass. The advantages of this method are that no addition of chemical flocculants is required and that similar cultivation conditions can be used for the flocculating microalgae as for the microalgae of interest that accumulate lipids. This method is as easy and effective as chemical flocculation which is applied at industrial scale, however in contrast it is sustainable and cost-effective as no costs are involved for pre-treatment of the biomass for oil extraction and for pre-treatment of the medium before it can be re-used.

Full text

Harvesting of microalgae by bio-flocculation
Sina Salim &Rouke Bosma &Marian H. Vermuë &
René H. Wijffels
Received: 22 June 2010 /Revised and accepted: 9 September 2010 / Published online: 28 September 2010
#The Author(s) 2010. This article is published with open access at Springerlink.com
Abstract The high-energy input for harvesting biomass
makes current commercial microalgal biodiesel production
economically unfeasible. A novel harvesting method is
presented as a cost and energy efficient alternative: the
bio-flocculation by using one flocculating microalga to
concentrate the non-flocculating microalga of interest.
Three flocculating microalgae, tested for harvesting of
microalgae from different habitats, improved the sedimen-
tation rate of the accompanying microalga and increased
the recovery of biomass. The advantages of this method are
that no addition of chemical flocculants is required and that
similar cultivation conditions can be used for the flocculat-
ing microalgae as for the microalgae of interest that
accumulate lipids. This method is as easy and effective as
chemical flocculation which is applied at industrial scale,
however in contrast it is sustainable and cost-effective as no
costs are involved for pre-treatment of the biomass for oil
extraction and for pre-treatment of the medium before it can
be re-used.
Keywords Harvesting .Microalgae .Bio-flocculation
Introduction
Oil-accumulating microalgae are a promising feedstock for
biodiesel production (Benemann et al. 1977; Lee et al.
2009). Commercial microalgal biodiesel production is not
economically feasible yet, mainly due to the high-energy
inputs required for water pumping, mixing and for harvest-
ing the microalgal biomass combined with large investment
costs (Schenk et al. 2008).
Harvesting in commercial microalgae production plants
is generally done by centrifugation. Different studies
showed a contribution of the costs for harvesting to more
than 30% of the total cost in case of algal production in
open ponds (Zittelli et al. 2006). These high costs can only
be justified in case of microalgal production for high value
products. For low-value bulk products, both the investment
as well as the operational costs should drastically decrease
to make commercial production feasible (Wijffels and
Barbosa 2010).
To minimize the energy consumption of harvesting
microalgae, an integrated approach is needed (Benemann
1997). Evaluation of several harvesting methods showed
that flocculation combined with flotation or sedimentation
and subsequent further dewatering by centrifugation or
filtration is the most promising cost and energy efficient
alternative (Schenk et al. 2008). During flocculation, the
dispersed microalgal cells aggregate and form larger
particles with higher sedimentation rate.
Flocculation can be induced in different ways. Induced
chemical flocculation using Zn
2+
,Al
3+
,Fe
3+
or other
chemical flocculants has been studied extensively
(McGarry 1970; Lee et al. 1998; Papazi et al. 2010) and
some of them are applied at industrial scale, especially in
wastewater treatment plants (De la Nouë et al. 1992).
Although this is an easy and effective method, this is not an
appropriate method for cheap and sustainable harvesting of
microalgae in large-scale microalgae production plants
because excess cationic flocculant needs to be removed
from the medium before it can be re-used and this leads to
extra operational costs (Schenk et al. 2008). Flocculation
can also be induced by changing the culture conditions by
S. Salim (*):R. Bosma :M. H. Vermuë :R. H. Wijffels
Bioprocess Engineering, Wageningen University,
P.O. Box 8129,
6700 EV Wageningen, The Netherlands
e-mail: sina.salim@wur.nl
URL: www.algae.wur.nl
J Appl Phycol (2011) 23:849–855
DOI 10.1007/s10811-010-9591-x

applying extreme pH, nutrient depletion, temperature
changes and changes of the level of dissolved O
2
. For
pre-harvesting of microalgae at large-scale these floccula-
tion methods are not preferred. Most of the latter methods
cannot be applied for controlled flocculation and they may
induce undesired changes in cell composition (Benemann
and Oswald 1996). All of them again require treatment of
the medium to be re-used (Schenk et al. 2008). The third
method that has been proposed for induced flocculation
of microalgae is biologically induced flocculation with
bacteria as has been applied successfully in wastewater
treatment (Lee et al. 2009). Bio-flocculation of microalgae
with bacteria, however, demands additional substrate as
well as an extra energy source for bacterial growth and
this will evoke undesirable bacterial contamination of
the algal production plant. Recently, the naturally floccu-
lating diatom Skeletonema was used to form flocs of
Nannochloropsis (Schenk et al. 2008). As diatoms have a
silica-based cell wall, they require different medium compo-
sition than most of microalgal strains used for biodiesel
production which leads to additional cultivation costs.
In this paper, bio-flocculation of a non-flocculating
microalga with another autoflocculating microalga has been
evaluated as a promising alternative effective method for
harvesting of microalgae. The presented bio-flocculation
method enables the harvesting of microalgae without
addition of chemical flocculants and allows for re-use of
the cultivation medium without any additional treatment.
Another advantage of this method in comparison with other
applied flocculating microorganisms (bacteria, diatoms) is
that it does not require different cultivation conditions and
therefore avoids additional costs and prevents undesired
contaminations. Furthermore the lipid content of the strains
used as the flocculating and non-flocculating microalgae in
this study is on average more than 25% of the dry weight
biomass (Table 1). The presence of the flocculating micro-
algae in the final biomass concentrate does thus not
interfere with further downstream processing of the lipids
into biodiesel. Unfortunately, the overall lower lipid
productivity of these flocculating microalgae makes them,
as such, less attractive for biodiesel production than the
faster growing non-flocculating microalgae (Griffiths and
Harrison 2008).
The bio-flocculation method will be compared with the
chemically induced flocculation, in terms of recovery
efficiency and time needed for sedimentation.
Materials and methods
Chlorella vulgaris (211-11b) and Scenedesmus obliquus
(276-3a) were obtained from University of Göttingen,
Germany (SAG), Neochloris oleoabundans (1,185) from
University of Texas, Austin, USA (UTEX), Tetraselmis
suecica (66/38) from SAMS, UK (CCAP) and Ankistro-
desmus falcatus (211) from the Center of Phycology,
Třeboň, Czech Republic (CCALA).
Culture conditions The marine medium contained NaCl
(27.00 g L
−1
), MgSO
4
·7H
2
O (6.60 g L
−1
), MgCl
2
·6H
2
O
(5.60 g L
−1
), CaCl
2
·2H
2
O (1.50 g L
−1
), KNO
3
(1.45 g L
−1
),
NaHCO
3
(0.04 g L
−1
), TRIS (hydroxymethyl) amino-
methane (3.94 g L
−1
), E D TA -Na
2
(95 μgL
−1
),
ZnSO
4
·7H
2
O(11μgL
−1
), CoCl
2
·6H
2
O(5μgL
−1
),
MnCl
2
·4H
2
O (90 μgL
−1
), Na
2
MoO
4
·2H
2
O (30 μgL
−1
)
and CuSO
4
·5H
2
O(5μgL
−1
) dissolved in demineralized
water. For the freshwater medium KNO
3
(3 g L
−1
),
NaH
2
PO
4
·2H
2
O (0.26 g L
−1
), KH
2
PO
4
(0.74 g L
−1
),
HEPES (2.38 g L
−1
), H
3
BO
3
(61.80 μgL
−1
), EDTA-Fe
(III)-Na, (0.11 g L
−1
), EDTA-Na
2
(37 mg L
−1
),
ZnSO
4
·7H
2
O (3.20 mg L
−1
), MnCl
2
·4H
2
O (13 mg L
−1
)
and CuSO
4
·5H
2
O (1.83 mg L
−1
) were added to demineral-
ized water. The pH of the solution was set at 6.8 using 4 M
HCl. One hundred milliliters of this medium was dispensed
into 300-mL Erlenmeyer flasks, sealed with cotton and an
aluminum cap and autoclaved for 20 min at 121°C. After
cooling the marine medium, K
2
HPO
4
(100 mg L
−1
),
KH
2
PO
4
(2 mg L
−1
), EDTA-Fe(III)-Na (1.36 mg L
−1
),vitamin
B
12
(1 μgL
−1
), D-biotin (1 μgL
−1
), and thiamine-HCl
(200 μgL
−1
) were added using a 0.2 μm non-pyrogenic
sterile filter (Sartorius Stedim Biotech, FR). For the freshwater
medium MgSO
4
·7H
2
O(0.4gL
−1
), CaCl
2
·2H
2
O(13mgL
−1
),
vitamin B
12
(1 μgL
−1
), D-biotin (1 μgL
−1
), and thiamine-
HCl (200 μgL
−1
) were added after cooling. The microalgae
were grown in a light and climate controlled shaking
incubator at 100 rpm and 25°C with a 2% CO
2
enriched
airflow (3 L⋅min
−1
), illuminated using fluorescent light
(50 μmol⋅photons m
−2
s
−1
) with a 16 h/8 h light/dark cycle.
Turbidity measurements Cell concentration was measured
as the optical density at 750 nm (OD
750
) with an Ultraspec
Table 1 Maximum and minimum reported lipid contents for the three
flocculating microalgal strains used in this study and for the two non-
flocculating microalgae
Strain Habitat Lipid content (% DW)
Flocculating microalga
A.falcatus Freshwater 28–37
S.obliquus Freshwater 21–42
T.suecica Marine 18–26
Non-flocculating microalga
C.vulgaris Freshwater 25–42
N.oleoabundans Marine 36–42
The data are adapted from Griffiths and Harrison (2008)
850 J Appl Phycol (2011) 23:849–855

2,000 spectrophotometer (Pharmacia Biotech Ltd. UK)
equipped with a temperature controlled carousel cell holder
with six positions. Demineralized water served as reference.
The microalgal samples were diluted in a 10× 10 × 45 mm
polystyrene cuvette (Sarstedt, DE) using filter-sterilized tap
water for the freshwater microalgae and with 0.46 mol⋅L
−1
NaCl solution (in demineralized water) for the marine
strains (similar ionic strength as the medium applied for the
marine strains) to achieve an OD
750
value below 1.
Sedimentation kinetics Samples of the microalgal suspen-
sions were taken and diluted in a cuvette. After mixing, the
suspension was left to settle at 27°C in the dark in a
spectrophotometer. The temperature and pH of all samples
were measured in the beginning and at the end of the
sedimentation period and they were constant respectively at
27°C and pH 7. During the settling period, turbidity of the
sample was measured at 750 nm at the same height in the
cuvette to determine the recovery. The microalgal recovery
(microalgal removal percentage) was calculated with:
recovery %ðÞ¼
OD750 t0
ðÞOD750 ðtÞ
OD750 t0
ðÞ 100 ð1Þ
where OD
750
(t
0
) is the turbidity of sample taken at time
zero and OD
750
(t) is the turbidity of the sample taken at
time t(Fig. 1). This was done for the suspension of non-
flocculating microalga with and without addition of the bio-
flocculating microalga. The sedimentation kinetics were
measured in cuvettes instead of in conventional jar tests
(Vandamme et al. 2010) or recently used cylindrical glass
tubes (Papazi et al. 2010). Similar to the conventional tests,
the recovery percentage is measured in the top part of the
cuvette, where individual cells and formed flocs indepen-
dently sink.
To compare different strains on their ability to be applied
as flocculating microalgae, the recovery efficiency is
defined as the recovery of the non-flocculating microalga
in the presence of the flocculating microalga divided by the
recovery of the non-flocculating microalga without floccu-
lating microalga present. The recovery efficiency (adapted
from Papazi et al. 2010 and Buelna et al. 1990) was
calculated with:
recovery efficiency %ðÞ¼ 1
ODa750ðtÞ
ODa750 t0
ðÞ
ODb750ðtÞ
ODb750 t0
ðÞ
2
43
5100 ð2Þ
where OD
a750
(t
0
)andOD
a750
(t) are the turbidities of
samples of non-flocculating microalga with flocculating
microalga taken at time zero and at time t, respectively.
OD
b750
(t
0
) is the turbidity of sample of non-flocculating
microalga taken at time zero and OD
b750
(t) is the turbidity
of the same sample taken at time t.
Three different flocculating microalgae were tested on
their ability to improve the recovery efficiency and the rate
of harvesting of the non-flocculating microalga. The
freshwater microalgae A.falcatus and S.obliquus were
used for harvesting of C.vulgaris. The marine microalga T.
suecica was used to harvest the non-flocculating marine
microalga N.oleoabundans. For each of the three tested
combinations of flocculating and the non-flocculating
microalga, four sedimentation experiments were performed:
(1) the flocculating microalga, (2) the non-flocculating
microalga, (3) the non-flocculating microalga with low
concentration of added flocculating microalga and (4) the
non-flocculating microalga with high concentration of
added flocculating microalga (Table 2). Each of these
experiments was performed in duplicate. At the end of
sedimentation experiment, samples were taken from the
bottom of cuvettes in order to make microscopic pictures of
the formed microalgal flocs.
Morphological analysis At the end of sedimentation
experiment, samples were taken from the bottom of
cuvettes in order to make microscopic pictures of the
formed flocs of the microalgal cells, using a C-3030 zoom
5-megapixel Olympus camera connected to a CK40
Olympus microscope with a SK20-SLP phase contrast filter
and a T6 objective (×40 magnification) and a NCWHK
18 L ocular lens (×10 magnification).
Results
Three different autoflocculating microalgae were identified;
the freshwater A.falcatus, and S.obliquus and the marine
Tetraselmis suecica (Fig. 2d, e, and f, respectively). The
freshwater microalgae were used to flocculate the strain C.
Fig. 1 Recovery of microalgal biomass and sedimentation kinetics
calculation. aSchematic overview of the microalgal sedimentation test
in time. bRecovery (%) of the microalgae from the suspension in time
J Appl Phycol (2011) 23:849–855 851

vulgaris as non-flocculating microalga (Fig. 2a and b),
while the marine microalgal strain was used to flocculate N.
oleoabundans (Fig. 2c). C. vulgaris and N.oleobundans
show both relatively high growth rates in comparison with
the autoflocculating microalgae, but all five microalgae are
reported to show relatively high lipid content (Table 1).
Microscopic analysis
Figure 2shows pictures of the non-flocculating microalgae
N.oleoabundans (Fig. 2c) and C.vulgaris (Fig. 2a and b).
The microalgae are present as single cells and no floc
formation is observed. In the sediments of all three
flocculating microalgae large flocs can be observed
(Fig. 2d, e, and f). If the three flocculating microalgae are
Tab l e 2 Optical densities (OD
750
(t
0
)) of flocculating and non-
flocculating microalgae added into the cuvettes for four combinations
of three experiments
Combination of flocculating and
non-flocculating microalgae
OD
750
(t
0
)
1* 2* 3* 4*
A.falcatus 0.7 0.7 0.4 0.0
C.vulgaris 0.0 0.9 0.9 0.9
S.obliquus 0.8 0.8 0.4 0.0
C.vulgaris 0.0 0.3 0.3 0.3
T.suecica 1.1 1.1 0.5 0.0
N.oleoabundans 0.0 0.2 0.2 0.2
1* the flocculating microalga, 2* the non-flocculating microalga; 3*
the non-flocculating microalga with low concentration of added
flocculating microalga; 4* the non-flocculating microalga with high
concentration of added flocculating microalga
Fig. 2 Microscopic picture of individual and flocculated microalgal
cells. The non-flocculating microalgae (aand bC.vulgaris and cN.
oleoabundans), the flocculating microalgae (dA.falcatus,eS.
obliquus and fT.suecica) and the flocs of the non-flocculating
microalgae after the addition of accompanying flocculating microalga
(gC.vulgaris with A.falcatus,hC.vulgaris with S.obliquus and iN.
oleoabundans with T.suecica). For more details on the morphological
analysis and sample preparation see “Materials and methods”
852 J Appl Phycol (2011) 23:849–855

added to the non-flocculating microalgae (Fig. 2g, h, and i),
the microscopic pictures show that the majority of the non-
flocculating microalgae are trapped in flocs formed by the
flocculating microalgae and almost no loose cells of non-
flocculating microalgae remain in the suspension after the
addition of flocculating microalgae. The comparison of the
pictures in Fig. 2a, b, and c, respectively, with Fig. 2g, h,
and iconfirms that the addition of flocculating microalgae
from different habitats (marine and freshwater) improves
the recovery of various non-flocculating microalgae.
Sedimentation kinetics of various flocculating
and non-flocculating microalgae
The sedimentation of the microalgal suspensions was
monitored for 8 h and the percentage of microalgal
recovery was determined over time. The sedimentation rate
of the microalgae in suspension was calculated by linear
regression of data in the curves of the recovery percentage
in time and use of the slope of the linear regression.
The initial sedimentation rates of the flocculating
microalgae measured over the first 2 h of the test are
higher than those of the non-flocculating microalgae
(Table 3). Mixing of the flocculating microalga with the
non-flocculating microalga increases the initial sedimen-
tation rate considerably. The large flocs formed by
flocculating microalgae seem to trap the non-flocculating
microalgae (Fig. 2g, h,andi) and sediment faster than
individual non-flocculating microalgal cells. Furthermore,
an increase in the ratio of the bio-flocculating microalga
and the non-flocculating microalga leads to higher
sedimentation rates. These observations again confirm that
the total recovery as well as the rate of sedimentation of
various non-flocculating microalgae improves upon addition
of different flocculating microalgae.
Efficiency of various flocculating microalgae
The improvement in the recovery of the non-flocculating
microalgae was evaluated for the three flocculating micro-
algae by calculation of the recovery efficiency percentage.
For calculation of the recovery efficiency percentage (Eq. 2
in “Materials and methods”), the average turbidity of
duplicate measurements was used. The standard deviation
in measured values for sedimentation rate and recovery
percentage for all tested samples was less than 3.5%. The
recovery efficiency percentage of three flocculating micro-
algae added at low and high concentration is presented in
Fig. 3.
All three flocculating microalgae show higher recovery
efficiency when they are applied at higher concentration,
although doubling of concentration of the flocculating
microalga does not necessarily result in two times higher
recovery efficiency of the non-flocculating microalga.
Discussion
The results show that addition of autoflocculating micro-
algae induce faster sedimentation of non-flocculating
microalgae and also increase the harvesting efficiency.
Similar positive effects on sedimentation rates and harvest-
ing efficiencies are observed with bio-flocculation of non-
flocculating microorganisms with bacteria (Lee et al. 2009).
In literature, adsorption of cationic polymers (Lewin 1956;
Tilton et al. 1972) excreted by the microorganisms is
proposed to explain the mechanism involved in bio-
flocculation. Polymer-induced flocculation can be divided
in two sub-mechanisms called bridging and patching
Table 3 Initial sedimentation rate
Combination of flocculating and
non-flocculating microalgae
Initial sedimentation rate
(% recovery⋅h
−1
)
1* 2* 3* 4*
A.falcatus and C.vulgaris 41.1 13.6 10.4 6.8
S.obliquus and C.vulgaris 37.0 20.4 18.7 10.2
T.suecica and N.oleoabundans 46.2 39.9 37.5 18.7
Details for the calculation of these initial sedimentation rates can be
found in the main text and “Materials and methods”
1* the flocculating microalga, 2* the non-flocculating microalga, 3*
the non-flocculating microalga with low concentration of added
flocculating microalga, 4* the non-flocculating microalga with high
concentration of added flocculating microalga
Fig. 3 Recovery efficiency percentage of different flocculating
microalgae at two different concentrations. filled square high
concentration T.suecica,empty square low concentration T.suecica,
filled diamond high concentration S.obliquus,empty diamond low
concentration S.obliquus,filled upright triangle high concentration A.
falcatus,empty upright triangle low concentration A.falcatus. The
standard deviation in measured values for sedimentation rate and
recovery percentage for all the tested samples was less than 3.5%.
Details for calculation of these recovery efficiency percentages can be
found in Materials and methods
J Appl Phycol (2011) 23:849–855 853

(Fig. 4). The positively charged polymers bind partly or
completely to microalgal cells. If the polymers bind partly,
the unoccupied part of the polymers can bind to other
microalgal cells, thereby bridging them and resulting in a
network of polymers and microalgal cells. If the polymers
bind the microalgal cells completely because they are too
short to bind others as well, they adsorb (patch) to the
surface and can create positive charges locally. These
charges attract other microalgal cells and also result in
flocculation of the cells.
Our microscopic observations suggest that bridging is
the mechanism behind the floc formation by A.falcatus
(Fig. 2d) as a large network of microalgal cells is formed.
Patching can be the mechanism behind the flocculation of
T.suecica (Fig. 2f) and S.obliquus (Fig. 2e) as they seem to
be connected more locally. Based on these observations,
our hypothesis is that the extracellular polysaccharides
excreted by A.falcatus itself bind partly to the surface of A.
falcatus and positively charged tails of these polysaccharides
can bind to the other A.falcatus cells. During the formation
of the flocs C.vulgaris cells are trapped in this large network
of A.falcatus cells (Fig. 5).
The recovery efficiencies and time needed for sedi-
mentation observed here using bio-flocculation are in the
same range as the recovery efficiencies found by Papazi
et al. (2010) applying chemically induced flocculation for
separation of the microalgal biomass. They showed a
recovery efficiency of 60% for harvesting Chlorella
minutissima by addition of 1 g L
−1
of Al
2
(SO
4
)
3
and
ZnCl
2
in, respectively, 1.5 and 6 h. The density of
microalgal culture (OD
750
)usedbyPapazietal.(2010)
was 2.4 which is comparable with the density of cultures
used in this study. Other studies using chemical floccula-
tion reported other concentrations and recovery efficien-
cies, e.g. Lee et al. (1998), and McGarry (1970)usedup
to, respectively, 300 and 125 mg L
−1
of Al
3+
. However,
the microalgal density of the samples used in these studies
is not mentioned and the recovery efficiencies are
calculated on a different way and therefore cannot be
compared with results of the current study.
Future perspectives of sustainable microalgal harvesting
We presented in this study that all three chosen flocculating
microalgae improved the recovery efficiency of the accom-
panying non-flocculating microalga. It can be concluded
that the bio-flocculation by using one flocculating micro-
alga for harvesting of another oil-accumulating microalga
can be applied as the controlled and reliable pre-
concentration step in harvesting of the oil-accumulating
microalgae, although large-scale experiments are still
needed to prove the feasibility and cost efficiency of this
method at industrial scale. Further, it was shown in this
study that different flocculating microalgal strains are
available for application of bio-flocculation in marine as
well as in freshwater environment. Using bio-flocculation
followed by sedimentation as the pre-concentration step
decreases the recovery time of the non-flocculating micro-
alga. The amount of flocculating microalgae used is still
relatively high in comparison with the non-flocculating
microalgae (Table 2). A decrease in the amount of
Fig. 5 Schematic view of the proposed mechanism involved in bio-
flocculation using A.falcatus as the flocculating microalga
Fig. 4 Schematic view of possible mechanisms involved in polymer-
induced flocculation; bridging and patching
854 J Appl Phycol (2011) 23:849–855

flocculating microalga by half did not show any major
effects on the recovery efficiency and time needed for
sedimentation of the non-flocculating microalga. This
indicates that this method is indeed promising and further
optimization of the ratio of the bio-flocculating microalga
and the non-flocculating microalga should be done to reveal
if large-scale utilization of this technique will indeed result
in considerable decrease of harvesting costs and energy.
To summarize, this harvesting method is as easy and
effective as chemically induced flocculation which is
applied at industrial scale, however in contrast to induced
chemical flocculation, this method is sustainable. Although
the cultivation of flocculating microalgae requires some
extra nutrients and energy, the flocculating microalgae do
not require an additional set of nutrients for cultivation in
comparison with the microalgae of interest. In the econom-
ical analysis of large-scale application of this promising
harvesting method the additional costs for a separate
cultivation system for cultivation of the flocculating micro-
alga should also be taken into account. In addition, the
flocculating microalgae accumulates lipids and no extra
operational and investment costs are involved for treatment
of the sediment (microalgal biomass) for further down-
stream processing towards biodiesel or for pre-treatment of
the medium before it can be re-used.
Acknowledgments This work was performed at Wetsus, Center Of
Excellence For Sustainable Water Technology. Wetsus is funded by
the ministry of economic affairs. The authors like to thank the
members of the theme ‘Algae’from Wetsus for the fruitful discussions
and especially the participating AF&F, DeAlg B. V., Delta, Dow
Chemicals, Eneco Energie, Essent, Friesland Foods, Hubert, Ingrepro,
Liandon, Neste Oil, Syngenta, and Unilever for their support.
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