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.

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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

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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 ~300-

500nm) 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.

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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 pre-

versus 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: 215-

220.

8. Overmann, J. (2005) Chemotaxis and behavioural physiology of not-yet-cultivated

microbes. Methods in Enzymology, 397.

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: ujaekel@mpi-bremen.de Tel. +49 421 2028 748