Ulrike Jaekel
Microbial Diversity 2010
Microbial degradation of chlorophyll
Abstract
Chlorophylls are abundant molecules in the environment. They are directly involved in
the light harvesting process of photosynthesis. Previous studies have shown that
chlorophylls do not accumulate in nature and are transformed by biological processes.
The biological transformations of chlorophyll through endogenous enzymes of plants
and algae have been studied in detail in the past. The microbial transformation of
chlorophylls in the presence and also in the absence of molecular oxygen has however
received little to no attention thus far. It was therefore the aim of this study to enrich for
aerobic and anaerobic microorganisms that could use chlorophyll as the sole substrate
for growth. An aerobic enrichment culture growing in minimal seawater medium with
chlorophyll in an inert hydrophobic carrier phase as the only carbon source was
obtained. Cells were found to be abundant both in the liquid and carrier phase of the
enrichment culture. The microorganisms in this culture showed a directed, chemotactic
movement towards the chlorophyll. Phylogenetic analysis revealed that the community
composition of the liquid phase and the carrier did not differ from each other
significantly.
1.
Introduction
Chlorophylls are abundant molecules in the environment, since they are directly involved
in the light harvesting process during photosynthesis which constitutes the major
process of primary production of biomass on the planet. Chlorophylls do not accumulate
in the environment, however the presence of their defunctinalization products, such as
chlorophyllides,
phaeophytin,
phaeophorbide,
pyrophaeophorbide
or
cycloalkanoporphyrins (CAPs) indicate that they are transformed in the biosphere. It was
found that the transformations of the major chlorophyll (a) to the major sedimentary
porphyrin (DPEP) require low temperatures and are most likely biologically mediated
(Bidigare et al, 1986, Ridout and Morris, 1988, Hule and Armsrong, 1990). Previous
studies have shown that chlorophyll (a, b) is transformed in plants and algae (see
Review by Matile and Hörtensteiner, 1999).
The removal of the central Mg-ion is mediated through a dechelatase, the cleavage of
the porphyrinring is perfomed by an oxygenase. The microbial degradation of chlorophyll
by microorganisms has for some reason not been studied in great detail thus far. Some
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studies report the disappearance of chlorophyll (a,b) in dead or live algae culture along
with an increase of bacterial cells in these cultures and the formation of chlorophyll
derivatives, indicating the transformation of chlorophylls in these cultures due to the
impact of microbial activity (Spooner et al., 1994a, 1995, Afi et al., 1996, Chen et al.
,2003). Furthermore, all of these studies have investigated the breakdown of chlorophyll
by microorganisms under oxic conditions. If oxygen is no longer present, as it is the case
in many environments (especially sediments), the question arises as to whether
chlorophyll can still be transformed by microorganisms which can use it a substrate for
growth? What would be the activation reactions involved (since cleavage of the
porphyrinring by oxygenases would seem unlikely)? It was therefore the aim of this
study to enrich for aerobic and anaerobic microorganisms that could use chlorophyll as
the sole substrate for growth.
2.
Materials and methods
2.1 Chlorophyll extraction
Chlorophyll as a substrate for growth in enrichment cultures was extracted from fresh
spinach (swope cantine) according to a protocol by Iriyama et al.,(1974), with the
following exception: acetone was used for chlorophyll extraction instead of methanol
because Khalyfa et al (1992) reported that using acetone instead of methanol increases
the overall yield. Furthermore, a plastic blender was unfortunately used instead of a
glass blender due to limitations in equipment. Also, the product of the second
water/dioxane
precipitation
was
not
further
purified
by
DEAE-Sepharose
chromatography due to the lack of available chromatographs and a suitable column
within the given time frame. For the above mentioned reasons it has to be taken into
consideration that the used extracted chlorophyll was most likely significantly
contaminated by traces of plastic from the blender and remaining carotenoids from the
spinach-chlorophyll extraction. The quality of the extracted chlorophyll was analyzed
with a spectrophotometer (Varian, Cary 50 Scan UV-Vis Spectrophotometer). After the
last precipitation step, the chlorophyll was dissolved in a minimal amount of acetone (a
few ml only). The acetone was subsequently evaporated under a constant N2 gas stream
at RT (Alu foil covered bottle to protect the light sensitive chlorophyll) until completely
dried.
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2.2 Set up of enrichment cultures with chlorophyll as the growth substrate
Enrichment cultures for chlorophyll degrading microorganisms were set-up from samples
taken from: 1. School Street Marsh (a water sample from close to a rotting seaweed), 2.
Sediment from a small park near School Street (underneath some growing grass and
fallen leaves) and 3. Guts from collected grazers from a freshwater pond in Falmouth.
The goal of the project was to enrich for microorganisms that could degrade chlorophyll
either under oxic or anoxic conditions. Therefore, sulphate, nitrate and iron (III) were
used as alternative electron acceptors. Table 1 shows an overview of all enrichment
cultures. Liquid cultures were set up in 50 ml serum bottles with 25 ml of either seawater
or freshwater minimal-, MOPS-buffered medium (lab manual). Extracted and dried
Chlorophyll was dissolved in the inert, nontoxic, hydrophobic carrier phase
Heptamethylnonane (Sigma) for a final concentration of ~1 mg/ml HMN. Per serum
bottle, 25 ml medium and 2 ml of Chlorophyll in HMN was added. For anaerobic liquid
cultures, anoxic minimal seawater medium and anoxic Chlorophyll in HMN were added
to the serum bottles in an anaerobic chamber (Coylab). All anoxic bottles were closed
with butyl stoppers and sealed with an alu crimp. Bottles were inoculated with 2 ml of
samples 1 or 3 (diluted gut extracts) or ~ 0.5 g of sample 2 (for anoxic enrichment
cultures, this was done in an anaerobic chamber). All liquid enrichment cultures were
incubated standing, at RT and in the dark (to prevent photooxidation of the chlorophyll).
For each set of liquid enrichment cultures (A-B, E-J) one abiotic control (Chlorophyll in
HMN, no inoculum) and one Chlorophyll-control (only HMN without Chlorophyll, with
inoculum) was included. Plate enrichment cultures were set up using minimal seawater
or freshwater agar plates (Agar Noble), which were overlayed with 3ml HMN dissolved in
Pentane. The Pentane was allowed to evaporate for ~30 minutes in a sterile hood.
Plates were inoculated by adding 2 ml of samples 1 or 3 and 2 ml of a slurry of sample 2
(10 g sediment in 50 ml freshwater or seawater medium). Plates were incubated upside
down after the sample had dried a bit into the agar at 30°C in the dark.
Figure 1. Set-up of culture bottes and plates for
enrichment cultures using chlorophyll as the sole
substrate for growth.
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Table 1. Overview of enrichment cultures for chlorophyll degrading microbes
Culture name
Liquid /Plate
Medium
Electron acceptor
Inoculm
A1
Liquid
Seawater
Oxygen
1
A2
Liquid
Seawater
Oxygen
2
B1
Liquid
Freshwater
Oxygen
1
B2
Liquid
Freshwater
Oxygen
2
B3
Liquid
Freshwater
Oxygen
3
C1
Plate
Seawater
Oxygen
1
C2
Plate
Seawater
Oxygen
2
D1
Plate
Freshwater
Oxygen
1
D2
Plate
Freshwater
Oxygen
2
D3
Plate
Freshwater
Oxygen
3
E1
Liquid
Seawater
Sulphate (28mM)
1
E2
Liquid
Seawater
Sulphate (28mM)
2
F1
Liquid
Freshwater
Sulphate (14mM)
1
F2
Liquid
Freshwater
Sulphate (14mM)
2
F3
Liquid
Freshwater
Sulphate (14mM)
3
G1
Liquid
Seawater
Nitrate (15mM)
1
G2
Liquid
Seawater
Nitrate (15mM)
2
H1
Liquid
Freshwater
Nitrate (15mM)
1
H2
Liquid
Freshwater
Nitrate (15mM)
2
H3
Liquid
Freshwater
Nitrate (15mM)
3
J1
Liquid
Seawater
Fe(III) (~3mM)
1
J2
Liquid
Seawater
Fe(III) (~3mM)
2
K1
Liquid
Freshwater
Fe(III) (~3mM)
1
K2
Liquid
Freshwater
Fe(III) (~3mM)
2
K3
Liquid
Freshwater
Fe(III) (~3mM)
3
5
2.3 Growth observations
Growth in liquid cultures was monitored by the development of turbidity in the liquid
phase (with settled sediment in cultures with inoculum from sample 2) and a
macroscopic change of the color or appearance of the chlorophyll/HMN phase. Samples
from the liquid phase and the Chlorophyll/HMN phase were looked at under the
microscope. Growth on plates was monitored by looking for the appearance of colonies
and associated clearing zones, indicating a breakdown of the chlorophyll (removal of the
central Mg-ion results in the loss of the specific green color). Positive growth resulted in
transfers of the cultures to either fresh liquid medium with overlayed chlorophyll/HMN or
a transfer onto plates with overlayed chlorophyll.
2.4 Chemotaxis experiment
A chemotaxis experiment with an aerobic liquid enrichment culture (culture A1) was
performed, which is similar to the setup published by Overmann (2005). A small
chamber was built using a glass slide and coverlids which are attached to each other
with molten paraffin (see fig. 2). Approximately 200 µl of liquid phase enrichment culture
were added to the chamber until it was completely filled. Three glass capillaries (0.1mm
thickness, Vitro Dynamics) were moved into the chamber, each filled with the following:
1. Chlorophyll in HMN, 2. HMN only and 3. Seawater medium.
Figure 2. Set-up of the chemotaxis chamber. The chamber is
filled with ~200µl undiluted enrichment culture A1. Capillary 1
contains Chlorophyll in HMN, Capillary 2 contains only HMN and
Capillary 3 contains seawater minimal medium.
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The movement of cells toward the three capillaries and the abundance of cells at each of
the capillaries was monitored using a microscope (x40 magnification). Strong movement
of cells towards and abundance of cells at capillary 1 but not capillary 2 or 3 after some
time was interpreted as positive chemotaxis towards chlorophyll.
2.5 Community analysis of positive enrichment cultures
The bacterial community of both the liquid phase and the Chlorophyll/HMN phase of a
positive enrichment culture (A1) was analyzed by constructing 16SrRNA clone libraries.
Genomic DNA was extracted from the two phases using the MoBio DNA extraction kit
for soil. PCR amplification of the 16SrRNA gene was done by using the universal
bacterial primers 8_forward and 1429_reverse. PCR reactions were set up by adding 0.5
µl of each primer (10µM) to 12.5 µl Promega 2x Master Mix, 9.5 µl PCR H2O, 2 µl of
gDNA were added as template for 16SrRNA gene amplification. PCR conditions were as
follows: initial denaturation at 94°C for 5 min, followed by 35 cycles of 94°C for 1 min,
45°C for 1 min and 72°C for 2 min, final eleongation was at 72°C for 10 min. Successful
amplification was verified by agarose gel electrophoresis (using a 1% agarose gel).
Bands were cut and purified using the Millipore DNA gel extraction kit. Purified bands
were cloned using the TOPO-TA PCR4 cloning kit for sequencing. Colonies were
picked, grown in overnight cultures and the inserts sequenced using the M13F primer.
Sequences were quality checked and analyzed using the RDP classifier.
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3.
Results
3.1 Extraction of chlorophyll
The extraction and partial purification of chlorophyll from spinach using a dioxane/water
precipitation resulted in a decrease of concentration of the carotenoids (peaks at ~300500nm) relative to chlorophyll a and b (peaks at ~650nm) after two times dioxane/water
precipitation (figure 3). The amount of chlorophyll obtained from the extraction was
approximately 1g/200g spinach.
Figure 3. Spectrophotometric analysis of spinach extracts before (A) and after the
second dioxane/water precipitation (B).
3.2 Enrichment cultures with chlorophyll as the growth substrate
Growth was observed on plates with cultures C1, C2 and C4 in the form of clearing
zones around colonies and bacteria attached to the chlorophyll (figure 4).
Figure 4. Growth observations on chlorophyll plates (A) versus no growth on
noninoculated chlorophyll plate (B). Microscopic image (x40 magnification) of
a colony picked from the grown chlorophyll plate before transferred to a fresh plate.
8
Growth was also observed in an aerobic liquid culture (A1). The liquid phase turned
turbid after ~ 5 days of incubation. Microscopic analysis revealed many motile bacteria
in the liquid phase of the enrichment culture, as well as some Protozoans. The
Microscopic analysis of the Chlorophyll/HMN phase showed that many bacteria were
attached to the carrier phase (Figure 6), sometime almost resembling a biofilm (A, D).
No turbidity was observed in the control bottle without chlorophyll/HMN but only HMN.
Figure 5. Microscopic observation (x100 magnification) of growth in the liquid
enrichment culture A1. Cells are frequently found to be attached to the chlorophyll
/HMN phase.
Analysis of the absorption spectrum of 20µl chlorophyll/HMN phase of the enrichment
culture A1 vs. the abiotic control showed that the chlorophyll peak (~650nm) had
decreased (data not shown). It should be investigated in the future if, using finer
analytical methods (HPLC, mass spectrometry) it is possible to detect the first metabolic
intermediates, i.e. the chlorophyll without the central Mg-ion (Pheophorbide) or the
cleaved porphyrinring can be detected. This could give indications about the involved
activation mechanisms used by the micoorgansims growing on chlorophyll under oxic
conditions. No turbidity was observed in any of the other enrichment cultures listed in
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table 1. This could be due to the short time course of the incubation period. It is possible
that it takes longer time for anaerobic bacteria which utilize sulphate, nitrate or iron (III)
as electro acceptor and chlorophyll as the carbon source to grow.
3.3 Chemotaxis experiment of microorganisms from an aerobic enrichment
culture towards chlorophyll
The chemotaxis experiment with the aerobic culture (A1) growing in seawater showed
that after incubation in the chemotaxis chamber for ~20 min many bacteria started
moving into the capillary which contained the chlorophyll/HMN phase (figure 6) (video on
data CD is called “oxic seawater 1a chlorophyll in HMN capillary), whereas almost no
bacteria had appeared in the capillary which contained only the HMN (video on data CD
is called “oxic seawater 1a chlorophyll HMN only capillary) or sweater. This finding
shows that there is a directed chemotaxis of bacteria from the enrichment culture
towards chlorophyll.
Figure 6. Chemotaxis assay investigation (x40 magnification). Most cells appeared in the capillary filled
with chlorophyll/HMN (A), whereas only few cells appeared in the capillary with only HMN (B).
3.4 Community analysis of an aerobic, chlorophyll degrading enrichment culture
The enrichment culture A1 (oxic, seawater, inoculate with water sample from School
Street Marsh) was further investigated regarding it’s bacterial community composition.
Since both in the liquid and the chlorophyll/HMN phase had been observed to harbor
many motile and carrier phase attached microorganisms, two 16SrRNA clone libraries
were constructed. This would allow seeing structural differences in the bacterial
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community composition between the liquid phase and the carrier phase which harbors
the actual chlorophyll by comparing the represented bacterial OTUs and their
phylogenetic affiliations. The sequencing of the clones for the liquid phase unfortunetyl
yielded only 21 good quality sequences, whereas for the chlorophyll/HMN phase 90
good quality sequences were obaine. This should be taken into consideration when
interpreting the results of the sequence analysis. The analysis of the 16SrRNA
sequences of both clone libraries (the data from the liquid phase is referred to as
“Chlorophyll 1” in the data CD, the data from the chlorophyll/HMN phase is referred to as
“Chlorophyll 2” in the data CD) using the RDP classifier revealed that most obtained
16SrRNA sequences represent phylotypes which belong to the Proteobacteria (figure 8).
Figure 7. Phyla distribution in clone libraries constructed from genomic DNA of the liquid (A,C) and
chlorophyll/HMN phase (B,D) of the oxic enrichment culture A1.
Breaking these further down, it can be seen that both phases show an
overrepresentation of OTUs which affiliate with the Gammaproteobacteria. The Liquid
phase seems to be more enriched in phylotypes belonging to the Alphaproteobacteria
than the Chlorophyll/HMN phase and some Deltaproteobacteria seem to be present as
well. An unweighted unifrac analyis (Lozupone et al, 2005) of the sequences from both
phases resulted in an average (1000 permutations) p-value of 0.4 within the Bonferroni
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correction, indicating that the phylogenetic composition of the two phases is not
significantly different. This can be interpreted by taking into consideration that many
bacteria appeared to be highly motile during the chemotaxis assay, showing a strong,
directed movement towards the chlorophyll/HMN phase. Therefore, it is possible that the
same bacteria are found both in the liquid phase and the chlorophyll/HMN phase,
moving towards the chlorophyll/HMN phase, feeding on it for a while, detaching, moving
towards the chlorophyll again and so on.
4. Discussion and conclusion
This study aimed to enrich for microorganisms that can use chlorophyll (a or b) as a sole
substrate for growth both in the presence and absence of molecular oxygen, since there
seems to be a gap in our understanding of the significance of microbial turnover of the
globally abundant chlorophyll. Based on the observations of this report, which indicate
that at least in the presence of oxygen chlorophyll seems to be utilized by bacteria,
future studies should focus on the activation mechanisms by which microorganisms can
activate the porphyrin ring structure- both in the presence and absence of molecular
oxygen. Furthermore, it would be of interest to study the phylotypes involved in the
turnover of chlorophyll and their abundance in the environment in order to understand
their significance in the global turnover of chlorophyll.
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References
1. Afi, L., Metzger, C., Largeau, C., Connan, J., Berkaloff, C and Rousseau, B. (1996)
Bacterial degradation of green microalgae : Incubation of Chlorella emersonii and
Chlorella vulgaris with Pseudomonas oleovorans and Flavobacterium aquatille. Org.
Geochem. 25 : 117-130
2. Chen, N., Bianchi, T. S. And Bland, J. M. (2003) Implications for the role of preversus post-depositional transformation of chlorophyll a in the Lower Missisippi River
and Luoisianna shelf, Mar. Chem. 40: 231-248
3. Spooner, N., Harvey, H.. R., Pearce, G.E.S., Eckhardt, C.B., Maxwell, J.R. (1994a)
Biological defunctionalization of chlorophyll in the aquatic environment II: Action of
endogenous algal enzymes and aerobic bacteria. Org. Geochem. 22: 773-780.
4. Matile, P. and Hörtensteiner, S. (1999) Chlorophyll degradation. Annu.Rev. Plant
Physiol. Plant Mol. Biol.. 50:67-95.
5. Szymczak-Zyla, M. and Kowalewska, G. (2008) The influence of microorganisms on
chloropjhyll a degradation in the marine environment. Limnol. Oceanogr. 53:851-862
6. Iriyama, K., Ogura, N. and Takamiya, A. (1074) A simple method for extraction and
partial purification of chlorophyll from plant material, using dioxane. J. Biochem.
76:901-904
7. Khalyfa, A. Kermasha, S. and Alli, I. (1992) Extraction, purification and
characterization of chlorophylls from spinach leaves. J. Agric.Food Chem. 40: 215220.
8. Overmann, J. (2005) Chemotaxis and behavioural physiology of not-yet-cultivated
microbes. Methods in Enzymology, 397.
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Contact and Acknowledgements
I would like to thank Elizabeth Wilbanks for helping me doing the
Chlorophyll extractions (just let me say- ice box!!!) and Ed Hall for
sampling. I have had so much fun working with you. I am furthermore
grateful to Karin Lemkau at WHOI who has also helped with the Chlorophyll
extraction by lending me her Rotorvap.
I would like to thank the Gordon and Betty Moore foundation for funding, as
well as the Max Planck Institute for Marine Microbiology in Bremen and my
supervisor Prof. Widdel for letting me take part in this course. Thanks to
Dan Buckley, Stephen Zinder, Rebekah Ward and all the TAs for organizing
this course. Finally, I would like to thank my entire class of 2010 for the
great time here. It was an incredible experience and I will never forget it!
Ulrike Jaekel
Max Planck Institute for Marine Microbiology
Celsiusstr. 1
D 28359 Bremen
Germany
Email: [email protected]
Tel. +49 421 2028 748