using lipid biomarkers to determine changes in community ... · glass amber sample bottles, 4 each,...

Using lipid biomarkers to determine changes in community

structure and ecological processes occurring in the meromictic

Sider’s Pond, Falmouth, MA.

Anika Aarons

12/19/2011

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Abstract

Lipid biomarkers were characterized at four depths within the water column (oxic zone,

oxycline, and two in the anoxic zone) at two sites in Sider’s Pond, Falmouth, MA. Samples were

also analyzed for chlorophyll, particulate organic C and N (POC/N) and stable isotopes (13

C

and 15

N). Surface sediment was also collected from each site for POC/N and lipids analyses.

Chlorophyll analysis included phaeophytin and bacteriochlorophyll a and chlorophyll a.

Chlorophyll a was indicative of purple sulfur bacteria (PSB), green sulfur bacteria (GSB) and

phytoplankton, while bacteriochlorophyll a distinguished between prokaryotic and eukaryotic

photosynthesizers with phaeophytin specific to PSB. All three photosynthetic pigments were

highest at the oxycline (5 m) indicating low productivity in the surface waters reflective of

reduced seasonal activity and high bacterial productivity at the oxycline. There was a large

difference in organic carbon isotopic fractionation between the oxic and anoxic zone. This

transition was further supported by the nitrogen isotopic fractionation indicating a shift from a

photoautotrophic dominated pathway to chemoautotrophic organisms. The POC and PON

followed similar trends of a minimum concentration within the oxic zone, increasing at the

oxycline, lower at 10 m and increasing to a maximum concentration at 12 m just above the

surface sediment. The lipid analysis involved cold ultrasonic organic extraction followed by

Folch extraction, transesterification and TMS-derivitization of the samples to produce samples

amenable for analysis by GCMS. Cholesterol and even-chained polyunsaturated fatty acids were

primarily indicative of phytoplankton and zooplankton in the oxic zone. Phytol corresponded

with the photosynthetic pigments to indicate the presence of purple and green sulfur bacteria at

the oxycline. Odd-chain saturated fatty acids (iso- and anteiso- C15 and C17), hopanoids and -

OH acids were indicative of bacteria in the anoxic zone. Stanols were indicative of microbial

transformation occurring throughout the water column and of early diagenesis occurring in the

sediment. Phytosterols, long chained alkanes and long, even-chained fatty acids were indicators

of allochthonous terrestrial inputs, such as plant waxes, that accumulated at depth and in the

sediment.

Key Words

Lipid biomarkers, zooplankton, phytoplankton, purple sulfur bacteria, sulfate reducing bacteria,

fatty acids, cholesterol, phytosterols, beta-hydroxy fatty acids, hopanoids, sterols, stanols,

alkanes

Introduction

Lipids are involved in many important biological processes such as energy storage and

cell membrane maintenance (Christie, 2003). Each species uses and constructs lipids in unique

ways that best suit their needs and functions. As a result, these lipids can be utilized as

biomarkers; compounds directly associated with certain ecological processes or set of organisms

in a particular ecosystem. In aquatic ecosystems, lipids in both the suspended and deposited

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organic matter tell us about their origins, how they’ve been formed and degraded, and even what

environmental physical conditions they have been exposed to, such as stratification. As particles

sink in the water column they can be transformed by zooplankton feeding and microbial and

chemical oxidation (Schefuβ et al., 2004). Prokaryotic bacteria and sulfate reducing bacteria

produce predomimantly more odd-chain length and branched saturated fatty acids (Table 1).

Eukaryotic phytoplankton produce even-chained and polyunsaturated fatty acids (PUFAs) which

are bio-accumulated in zooplankton. Cholesterol is a zooplankton biomarker. As unsaturated

compounds are degraded and processed by other organisms, they can become more saturated.

Bacteria also produce hopanoids and -OH acids and the phototrophic species also produce

phytol.

When at the sediment-water interface, 30-99% of organic matter is remineralized during

diagenesis through microbially-mediated processes. The degradation of the organic matter in the

sediments is controlled by physical and chemical environmental conditions, such as salinity and

electron acceptor concentrations, and its quality or the nutritional value it provides to benthic

organisms. These factors help shape the redox chemistry of the ecosystem and how well organic

matter is preserved in the sediment (Canuel and Martens, 1996).

My aquatic ecosystem of interest is Sider’s Pond, Falmouth, MA. It is often characterized

by three stratified layers of different salinities and an oxic zone extending to 4-6m depths.

Consequently, its community structure is dominated by phyto- and zooplankton in the oxic zone,

purple and green sulfur bacteria at the oxycline and sulfate reducing bacteria in the anoxic zone.

I therefore hypothesized that I will observe different biomarkers in each of these zones indicative

of these shifts in community structure. Some of the biomarkers that I expected to see were

phytosterols from phytoplankton, cholesterol from zooplankton, and polyunsaturated fatty acids

(PUFAs) from both phyto- and zooplankton. C15-C19 n-alkanes (especially C17) and phytols were

expected from some photosynthetic bacteria such as purple and green sulfur bacteria (PSB and

GSB, respectively), and -OH acids, hopanoids and short-chain (< 20 carbons) fatty acids from

sulfate reducing bacteria (SRB) (Canuel et al., 1995; Gossens et al., 1989; Huang et al., 2004;

Pearsons et al., 2007). I also expected to see high concentrations of terrestrial plant material in

the sediment due to its recalcitrant nature, such as long (>20 C) alkanes and long even chain fatty

acids from leaf waxes, and a low free to bound lipids ratio.

2. Methods

2.1 Location

The site that I investigated was Sider’s Pond, a meromictic pond ecosystem that is

groundwater fed but receives occasional saltwater input from a small channel that connects to the

Vineyard Sound at very high tides such as storm surges. The saltwater influx sinks rapidly below

the fresh groundwater as it has a higher density and this density difference creates stratification

in the pond. The difference in densities is enough that the water column does not mix and

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therefore oxygen is only observed in the first 5.5 meters of relatively freshwater. The oxycline is

located at around 5.5 m. The chemistry of the pond is heavily influenced by chemoautotrophs

such as purple and green sulfur bacteria at the oxycline and sulfate reducing bacteria below in the

anoxic zone. Because the pond does not turn over and there is limited oxygen availability, no fish

are to be found in the pond.

2.2 Sampling

Sarah Nalven and I collected data from Sider’s Pond on November 14th

, 2011 to attain a

physical profile and to collect samples for organic and particulate organic carbon and nitrogen

(POC/N), and chlorophyll analyses. Sarah collected water samples for sulfide and sulfate, and

DNA analyses. Before heading into the field I placed two 8oz. glass jars, and eight 4L and 1L

glass amber sample bottles, 4 each, into a cooler and partitioned the bottles using cardboard to

prevent them from knocking against each other. The two glass jars were intended for the surface

sediment samples, the 4L bottles for water to be analyzed for organic lipids, POC/PON, and bulk

isotope analyses, and the amber 1L bottles for water for chlorophyll analyses. I also prepared an

insulated bag containing two of these 1L bottles that couldn’t fit into the cooler. Ice packs were

inserted around the bottles to keep the water samples cool in the field until return to the lab. All

of the glass jars had been previously cleaned in a Nochromix inorganic oxidizer solution in

concentrated sulfuric acid and combusted overnight at 380°C. The 4L glass bottles used to

collect water samples for organic biomarker analysis were empty solvent bottles previous

containing Optima grade organic solvents.

It was a very windy day with easterly winds creating small waves on the surface of the

water. We measured the total depth of Site 1 to be approximately 12.3m using a depth finder.

The Hydrolab was used to get a physical profile of the water column – temperature, PAR, pH,

salinity, conductivity, DO (mg/L), and DO% – at every meter except in the oxycline where it

was taken every half meter (4-6m). I used this information to then determine where to take water

samples; one sample in the oxic zone, one at the oxycline, and two in the anoxic zone.

Consequently, site 1 water samples (4L and 1L) were taken at 2, 5.5 (0.02 mg/L DO), 10.25 and

12m depths using a peristaltic pump.

I had a replicate Site 2 with a very similar physical profile and a slightly shallower total

depth of approximately 11.7m. The Hydrolab was used to take the same physical parameters

from 3 to 5.5m. I collected water samples at 2m, 5.5m, 10m, and 11.25m.

The Site 1 surface sediment sample was collected by dragging an anchor along the

bottom of the pond transferring the sediment from the rope knot attached to the anchor into the

glass jar using a spatula while the Site 2 sediment sample was collected by disturbing the

sediment bed with the tubing and then pumping it up into the 8 oz. jar. This produced a black

slurry, which therefore may have made it a sample of the sediment-water interface. The sediment

was separated from the water in the laboratory using centrifugation.

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2.3 Laboratory Methods

The 4L bottles and glass jars were immediately placed into a walk-in refrigerator (4°C)

until further analysis. The chlorophyll a samples were vacuum filtered onto combusted 47mm

glass fiber filters (GF/Fs). Each filter was carefully folded in half and wrapped in labeled foil.

They were then put into individual Ziploc bags and put into the -30°C freezer until an analysis

for chlorophyll a, phaeophytin and bacteriochlorophyll a was carried out eight days later using a

modified Lorenzen (1967) method. Sarah Nalven carried out sulfide and sulfate analyses using

modified Gilboa-Garber (1971) and ion chromatography, respectively.

On the 15th

of November, each of the 4L samples were vacuum-filtered through three

combusted 47mm GF/Fs (for lipids) and two combusted 25mm GF/Fs (for POC/N and stable

isotope analysis, 13

C and 15

N) until the filters were fully loaded with sample particulate

material. The samples were shaken vigorously between each volume addition to assure

homogenous resuspension of particulates. The three 47mm filters from each sample were

combined into one 40 mL glass vial and immersed in 2:1 chloroform:methanol organic solvent

(Fisher® Optima grade). These vials were then sealed using Teflon tape and placed into the -

30°C freezer overnight. The 25mm GF/Fs were acidified to remove inorganic C by placing them

on clean petri-dishes in a hydrochloric acid fuming chamber overnight. The sediment samples

were transferred to acid-cleaned 250mL LDPE bottles that were uncapped and placed into large

glass jars attached to the freeze dryer. A combusted 47mm GF/F filter was secured on the top of

each open LDPE bottle using perforated combusted aluminum foil to prevent the risk of fine

material escaping due to the vacuum. The freeze dryer condenser attained -100°C and a vacuum

of 0 mtorr within 3 hours. The samples were left on the freeze dryer for 48 hours to ensure

complete dryness.

The following day an extraction blank was prepared by suspending three combusted

47mm GF/Fs in 2:1 chloroform:methanol in a clean, combusted 40mL glass vial. This blank was

placed with the other glass vials that were removed from the freezer. A solvent rinsed 20µL

Hamilton syringe was used to add 15µL of a previously prepared trap standard solution (TRAP

STD 8/07 containing four internal standard compounds representing the major biomarker classes

expected: 362.1 ng/µL 21:0 Fatty Alcohol, 348.4 ng/µL 23:0 Fatty Acid, 354.0 ng/µL 5

cholestane, and 357.1 ng/µL 36 n-alkane) to all pelagic samples and the extraction blank. The

vials were then rewrapped with Teflon tape and ultrasonicated (100W, 5 min.) in a cuphorn filled

with recirculating cold ethylene glycol to prevent thermal degradation of the organic compounds

during extraction. The exterior of the vials was then cleaned with acetone to remove any

remaining ethylene glycol coolant and returned to the -30°C freezer. The 25mm GF/Fs were

transferred from the fume hood to petri slides and later folded in foil and pelletized for

particulate organic carbon and nitrogen (POC/N) analysis (quantification and stable isotopes)

using a ANCA-SL EA-GC interfaced with a Europa CF-IRMS.

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An aliquot of the freeze-dried sediment samples (3-5g) was immersed in 2:1

chloroform:methanol organic solvent, ultrasonicated (100W) twice for ten minutes each, and

centrifuged for 15 minutes.

For both the pelagic and surface sediment samples, lipids were extracted using a modified

Folch Extraction process. The samples were transferred to combusted, solvent rinsed fritted

funnels using combusted, solvent rinsed Pasteur pipettes. The sides of the sample vials were

rinsed twice with 2 mL of the 2:1 chloroform:methanol solvent and transferred to the funnels

after each rinse. The filters were added and pressed down onto the fritted glass using clean filter

forceps to ensure that most of the solution was extracted. In the case of the sediment samples,

special care was taken to ensure that no sediment was transferred. The samples filtered through

the fritted funnels, with the aid of a vacuum, into 60 mL solvent rinsed separatory funnels and 10

mL of 0.88% aqueous KCl added to each. The funnels were then stoppered and shaken for a

minute, releasing the pressure briefly at a point during the process, and then returned to the stand

for the phases to separate. The lower organic layer of each sample was transferred to a clean,

labeled 50 mL pear-shaped flask. The process was repeated with the remaining aqueous layer in

the funnel using 10 mL chloroform to extract any organics left behind. Each of the pear-shaped

flasks were placed onto the rotoevaporator (25ºC, 25”Hg) until just-dry and then resuspended in

500 µL chloroform. These lipid extracts were then filtered through clean Na2SO4 minicolumns

(~2 inches of combusted (June 15,2011) and desiccated Na2SO4 in a 5.5 inch Pasteur pipette

plugged with glass wool and cleaned with 2 mL hexane) into combusted, solvent-rinsed 13 mm

Pyrex vials to remove any residual moisture. The pear-shaped flasks were rinsed with 500 µL

chloroform three times and run through the columns to be transferred to the 13 mm vials.

These samples were then evaporated in the Savant for 30 minutes (1-2 torr) followed by

resuspension in 500 µL toluene and lipid transesterification under N2 using 10% methanolic HCl

(550C, 12 hours) to breakup to complex lipids into component classes and esterify the free fatty

acids. The transesterified samples were hexane extracted (1.5 mL of aqueous 5% NaCl added

and extracted with hexane (2mL, then 2x 1mL). The hexane extracted was dried passing through

Na2SO4 minicolumns into clean labeled centrifuge tubes, evaporated to just-dry for 30min using

the Savant vapor trap (at 1-2 torr vacuum), resuspended in dichloromethane (DCM) and

transferred to labeled 2mL vials.

Samples were TMS-derivitized for GCMS analysis. Samples were evaporated under N2,

and resuspended in 5 drops (~50 µL) of dichloromethane (DCM). Vials were tilted and rotated

until all the sample had dissolved. The samples were also rubbed to create warmth to dissolve

and 36 alkane that may have precipitated out during evaporation. The extract in DCM was

transferred to clean labeled V-vial Agilent autosample vials and evaporated with N2. A drop of

acetone (~10 µL) was added to each vial and evaporated to ensure no residual moisture before

TMS-derivization. Equal amounts of N,0-Bis (trimethylsilyl) trifluoroacetamide with

trimethylchlorosilane catalyst (BSTFA+1%TMCS, Sigma-Aldrich) and pyridine (1:1 v:v) were

added to the dry sample to silyate alcohol groups. Vials were capped under N2 and the reacted in

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the oven (550C, 1 hr). The samples evaporated under N2 and then resuspended in

dichloromethanol (DCM) for GC-MS analysis. After an initial screening run, an additional C36

alkane external standard (58.57 µg) was added to each of the sediment samples before

quantitative GCMS analysis since the extractable lipids concentrations were determined to be

much higher than initially expected. The GC-MS was an Agilent 7890A gas chromatograph

interfaced to an Agilent 5975C MSD with a triple axis detector. The GC-MS column was a 60m

CP Sil 5CB and the carrier gas was ultra high purity helium. The gas chromatograph temperature

program was 50ºC for two mins increasing to 150ºC at a rate of 10ºC/min and then to 320ºC

(4ºC/min) with a 40min hold. The GCMS has dual detectors: part of the sample stream is

diverted to a flame ionization detector (FID) for peak quantitation and the rest enters the mass

spectrometer for peak identification from the characteristic mass spectra.

2.4 Statistics

All the results reported are averages of the two sites and the standard deviation between the two

with the exception of the physical profile, chlorophyll a, sulfate and sulfide data that only reflect

Site 1. Additionally, if a compound was only measured at one site then its concentration was

used as the average and the standard deviation set to zero. Standard deviation of a compound

class was calculated as a sum of the standard deviations of each compound within that class.

3. Results

3.1 Physical profile

Sider’s Pond had two distinct haloclines (4 and 8 m) and an oxycline at 5.5m depth (Fig.

1). The temperature was fairly constant at 11.3°C in the first three meters of the pond and then

began increasing until 6 m depth, 17.2°C. Below 6 m the temperature steadily decreased to

below the surface temperature, 11.04, at 12.2 m. The euphotic zone ends at around 3m depth but

the light intensity doesn’t drop below 20 µE m-2

s-1

until 5 m depth and is >5 µE m-2

s-1

throughout the water column (Fig. 2). The pH pond of the pond steadily decreased from 7.53 at

the surface to the most acidic point in the pond at 5 m with a pH of 6.76 (Fig. 3). Below 5 m the

pH steadily increased to 7.11 at 12.2 m.

3.2 Sulfur chemistry

Sulfate was present throughout the water column; 1126-1603 µM in the surface then

steadily increased from 3 m to 7 m (Fig. 4). Below 7m there was great fluctuation in the sulfate

concentration. Sulfide only appeared below 6 m, and steadily increased with depth (Fig. 5).

3.3 Chlorophyll

Chlorophyll a was lowest at 2 m (5.3 µg/L), peaked at the oxycline (14.3 µg/L) and then

steadily decreased with depth, 9.2-6.9 µg/L (Fig. 6). Phaeophytin had a similar trend but the

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concentration slightly rebounded at 12 m (Fig. 7). The same trend was observed in

bacteriochlorophyll a except it was not present at 2m.

3.4 POC/N, TEL and Stable isotopes

The particulate organic C and N (POC/N) concentrations were lowest at 2 m and highest

at 12 m (Fig. 8, 9). However, these concentrations did not steadily increase with depth as there

was a significant decrease at 10 m. The isotopic fractionation of particulate organic carbon

changed from -35‰ in the oxic zone to -29-30‰ at and below the oxycline (Fig. 10). The

nitrogen fractionation was highest at 2 m (7‰) and declined dramatically at the oxycline (2‰)

then increased slightly in the anoxic zone (Fig. 11). The total extractable lipids (TEL)

concentration followed the same pattern as the total organic C concentration but had the

maximum concentration at the oxycline (201.25µg/L) instead of at 12 m (Fig. 12). When

normalized to organic C, the TEL is still highest at the oxycline but steadily decreases deeper in

the water column to a concentration much lower than at 2 m (Fig. 13).

3.5 Fatty Acids

The total fatty acids (TFA) concentration followed the same trend as the TEL – highest at

the oxycline (81 mg/L) – however, when normalized it was most prevalent in the oxic surface

water and declined with increasing depth (Fig. 14, 15). A general shift was observed from

primarily even-chain saturated fatty acids in the oxic zone to an increasing presence of odd-chain

saturated fatty acids in the anoxic zone (Fig. 16). The PUFAs composed of approximately 40%

of the TFA in the oxic zone (Fig. 17). The TFA saturation trend was a shift from unsaturated

fatty acids as most prevalent in the oxic zone to saturated fatty acids at depth. In the oxic zone all

of the unsaturated fatty acids (PUFAs and mono-unsaturated) were even chained (Table 2) and

86% of the saturated fatty acids were even chained. The even chained unsaturated fatty acids

decreased with depth while the odd chain saturated fatty acids increased with depth. The average

TFA sediment concentrations were 2443.27 µg/g dry weight and 17.45 µg/mg Organic C.

3.6 Phytosterols, cholesterol and phytol

Phytosterols and cholesterol concentrations increased with depth, although cholesterol

was slightly less at the oxycline (174.2 ng/L) than in the oxic zone, 250.3 ng/L (Fig. 18, 19).

Phytol was lowest in the oxic zone (52.8 ng/L), spiked at the oxycline (393.2 ng/L) and then

decreased in the anoxic zone, 252.3 - 176.7 ng/L. When normalized to the POC measured at each

depth, cholesterol was highest at 2m, lowest at the oxycline and slightly more elevated in the

anoxic zone (Fig. 20). Phytosterols were also lowest at the oxycline (2281 ng/L) but highest at 10

m (5635 ng/L) instead (Fig. 21). Phytol followed the same trend as it did pre-normalization

except that the ratio at 10 m was only slightly lower than the ratio at the oxycline. Within the

phytosterols, campesterol, stigmasterol, sitosterol, brassicasterol, and its 5 stanol increased with

depth (Fig. 22, 23). Cholesta-5,22E-dien-3-ol was only observed in the surface water samples.

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3.7 Alkanes

The total alkane concentration fluctuated throughout the water column; it was lowest at 2

m (108.1 ng/L), higher at the oxycline (220.6 ng/L) lower at 10 m (246 ng/L), and highest at 12

m, 347.6 ng/L (Fig. 24). When normalized to POC the total alkane concentration followed the

same pattern, fluctuating between 0.1 and 0.2 µg/µg Organic C (Fig. 25). Hexadecane and C17

alkane were the only short chain (<20 C) alkanes extracted from the pelagic samples (Table 3).

The concentrations of hexadecane and the C17 alkane increased with depth with hexadecane only

occurring at 10 and 12 m. Long chain (>20 C) alkanes were present throughout the water column

at low concentrations and peaked at 5.5m (Table 4). However, the largest concentration of long

chain alkanes was in the sediment where it was much higher than the short chain alkanes.

3.8 Hopanoids and -OH fatty acids

Hopanoids and -OH fatty acids had similar positive trends between concentration and

depth although the total -OH acids concentrations were an order of magnitude higher than that

of the hopanoids (Fig. 26, 27). The sediment concentrations were 95.21 and 119.23 µg/g dry

weight, respectively. When normalized to organic C, both compounds once again had similar

trends with the highest concentration at 10m and the lowest concentrations at the oxycline and in

the oxic zone (Fig. 28, 29). The normalized sediment concentrations were 0.68 and 0.86 µg/mg

Organic C respectively.

3.9 Sterols and Stanols

The concentration of sterols steadily increased with depth (2.19 – 11.71 µg/L) while the

stanols were lower at the oxycline (0.45 µg/L) that in the surface water (1.76 µg/L) and steadily

increased in the anoxic zone. 1.58 – 2.45 µg/L (Fig. 30). When normalized to POC both stanols

and sterols were lowest at the oxycline (Fig. 31). Stanols were most prevalent in the surface

waters while sterols were most prevalent at 10 m. The sterols concentration was consistently

higher than the stanols throughout the water column. The concentration of sterols was also higher

than that of stanols in the sediment at 642 and 48.6 µg/g dry weight respectively.

Discussion

The replicate sites showed an analogous physical profile that was reflected in the

similarity between their lipid compositions. There were distinct differences between the zones

that I sampled. The 10 and 12 m samples within the anoxic zone were the most comparable in

composition. The observed differences between the zones were depictive of the changes in

pelagic community structure and environmental conditions.

The low particulate organic carbon (POC) and chlorophyll a concentrations at 2 m were

indicative of low productivity in the oxic zone which is likely due to the beginning of the winter

season (Chen et al., 2001; Pimenov et al., 2008). Evidence of low winter productivity was

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presented in the low temperatures recorded at the site in addition to POC and stable isotope

measurements. The large accumulation of POC at 12 m depth is indicative of a downward

particle flux likely due to a recent seasonal die off of organisms in the upper waters.

Furthermore, the isotopic lightness of the POC in the oxic zone could be due to the aerobic

destruction of organic matter from phytoplankton. On the other hand, it could also be indicative

of terrestrial input from runoff following a recent rain event (Pimenov et al., 2008).

This very light 13

C signal in the oxic zone became significantly more enriched at the

oxycline suggesting that there is a large active bacterial community thriving in the pond’s

warmer waters. Evidence of this is provided in a similar study conducted by Pimenov et al.

(2008) in the meromictic Lake Mogil’noe, Russia. A similar trend in organic carbon

fractionation was observed with the heaviest isotopic composition in the anoxygenic

phototrophic bacteria (APB) development zone at the oxycline. The isotopic fractionation above

and below the oxycline was 2-6‰ lighter, the same trend that I observed in Sider’s Pond.

Pimenov et al. (2008) took readings immediately following the melting of the ice covering the

lake and a time of low photosynthetic productivity. In Sider’s Pond, the decline in POC from 5.5

to 10 m depth indicated movement away from the photosythetically productive zones and of

organic matter mineralization by sulfate reducers.

The difference in nitrogen fractionation between the oxic and anoxic zones showed a

clear transition between two different communities living in very different environmental

conditions. The 15

N signal can indicate the trophic level at which an organism is feeding in

addition to their environmental conditions. For example, an increased availability of ammonium

from organic matter mineralization may cause chemoautotrophic bacteria to preferentially

assimilate 14

N over 15

N resulting in higher isotope fractionation and a lighter, more depleted

15N value (Roach et al., 2011). The enrichment in 15

N concomitant with a decrease in PON at

10 m reflects fractionation occurring in the remineralization of N (Montoya et al., 1992). A

similar phenomenon is likely occurring in the sediment that has accumulated a lot of organic

matter over time from internal and external inputs and results in a15

N value comparable to that

of the oxic zone.

The total extractable lipids (TEL), as a component of POC, logically follow the same

concentration trend as POC. However, below the oxycline the TEL composed a very small

portion of the total organic C relative to the rest of the water column. This may depict a higher

concentration of non-lipid organic C present in the higher salinity, primarily ocean-fed layer

compared with the fresher groundwater-fed surface water.

The most abundant TEL compound class was the fatty acids. Branched and odd-chained

fatty acids are characteristic of bacteria; especially the short-chained iso- and anteiso-C15 and C17

saturated fatty acids indicating the presence of sulfate reducing bacteria (Canuel et al., 1995;

Pearson et al., 2007; Rotani and Volkman, 2005). On the other hand, even-chained unsaturated

fatty acids – such as C20 and C22 polyunsatured and C16:1 fatty acids – and even-chained

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saturated C14:0 and C16:0 are indicative of zoo- and phytoplankton species (Canuel et al., 1995;

Hama 1991). Even--chained saturated C14:0 and C16:0 fatty acids and C16:1 are synthesized by

many organisms but are more indicative of these species in this type of system and are also less

abundant in animal lipids (Canuel et al., 1995; Christie, 2003). Furthermore, the even chained

unsaturated and saturated fatty acids dominated the TFA in the oxic zone positively indicating

the presence of these species. The short-chained iso- and anteiso-C15 and C17 saturated fatty acids

increased with depth indicating that SRB was the dominant species in the anoxic zone as

expected. The increasing abundance of SRB biomarkers with depth corresponded to the observed

sulfur chemistry of Sider’s Pond. The fluctuations in the sulfate concentrations at depth could be

SRB converting SO42-

to S- as the sulfide concentrations steadily increase with depth in the pond.

The phytosterols observed were cholesta-5,22E-dien-3-ol, campesterol, stigmasterol, -

sitosterol, brassicasterol, and its 5 stanol. Campesterol, stigmasterol and -sitosterol are the

most abundant phytosterols found in vascular plants, algae, phytoplankton and some

cyanobacteria (Ali et al., 2009; Pearson et al., 2007). The accumulation of the phytosterols at

depth, out of the euphotic zone, with the exception of cholesta-5,22E-dien-3-ol, is therefore

primarily indicative of allochthonous plant material and/or phytoplankton debris sinking through

the water column. This evidence was further supported by the high concentrations of

campesterol, stigmasterol and -sitosterol observed in the sediment. Of the phytosterols,

cholesta-5,22E-dien-3-ol was only observed in the oxic zone sample. It is a commonly found

in some phytoplankton species and also in zooplankton carcasses, molts and feces (Colombo et

al., 1996). This may appear to be evidence of the zooplankton presence in the oxic zone and

possibly of a recent flux of plankton debris from the overlying surface waters, as I previously

theorized. Cholesterol, which is also an indicator of zooplankton tissues and fecal material, was

observed throughout the water column, indicating that cholesta-5,22E-dien-3-ol is not a reliable

measure of zooplankton. (The absence of cholesta-5,22E-dien-3-ol below the oxic zone may be

an artifact as it elutes from the GC column closely after brassicasterol. This could explain why

these two compounds were never recorded together in any samples.)

Cholesterol is found in high abundance in animals, such as zooplankton, and their feces.

It is produced by zooplankton through the metabolic processing of consumed phytosterols

(Pearson et al., 2007; Rotani and Volkman, 2005). Cholesterol also occurs as a minor sterol

component in some phytoplanktonic species such as diatoms and also in microflagellates such as

dinoflagellates which are found in both marine and fresh water systems (Schefuβ et al., 2004).

Cholesterol was found to be highest at the surface where I would expect to find these eukaryotic

zooplanktonic species.

The limit of the euphotic zone is above the oxycline at 5.5 m but purple sulfur bacteria

can photosynthesize under very low light conditions (5-10µE m-2

s-1

) and only tolerate very low

oxygen or anoxic conditions (Franks and Stolz, 2009; Stal et al., 1985). Green sulfur bacteria

have also demonstrated the ability to thrive under such low light conditions; brown-pigmented

12

cultures showed optimal growth within the light intensity range of 5-50 µE m-2

s-1

(Chen et al.,

2001). The minimum of both PSB and SRB’s light ranges were not recorded in Sider’s Pond.

Phytol is formed from the base hydrolysis of bacterial chlorophyll a by purple sulfur bacteria in

the genera Rhodobacter, Halochromatium and/or Thiohalocapsa and derived from the side chain

of chlorophyll a of phytoplankton (Pearson et al., 2007; Rotani and Volkman, 2005; Shioi and

Sasa, 1984). Phytol and chlorophyll a followed the same trend to obtain maximum

concentrations at the oxycline (5.5 m). The other pigment biomarkers, phaeophytin and

bacteriochlorophyll a, concentrations were also highest at 5.5 m, and sulfide concentrations were

undetectable until 7 m, further punctuating the presence of purple and green sulfur bacteria

within this transitional zone (5-7 m). The overall concentration of phytol was uncharacteristically

low which suggests that it may have been transformed by sulfate reducing bacteria to its

degradation compounds, phytadienes and phytane, which may have gone unidentified (Goossens

et al., 1989; Grimalt et al., 1992).

Short chain alkanes, especially n-C17 alkane, are a biomarker for algae and photosynthetic

bacteria as opposed to long chained terrestrial plant inputs (Huang et al., 2004; Meyers, 2003).

However, the concentration of n-C17 alkane increased with depth suggesting that it is not an ideal

biomarker for photosynthetic bacteria that are expected to be primarily located at the shallower

oxycline (Franks and Stolz, 2009). The other short chain alkane observed was hexadecane, which

a known resource for sulfate reducing bacteria and has been used in cultures to promote SRB

growth (Aeckersberg et al., 1991). This compound may not have been detectable until the anoxic

zone because of rapid bacterial consumption of the compound at lower concentrations higher in

the water column. Downward particle flux may have allowed it to accumulate to a detectable

concentration that was large enough to exceed the rate of SRB consumption. The hexadecane

concentrations within the anoxic zone are still relatively low. The absence of other short-chained

n-alkanes throughout the water column was depictive of rapid biodegradation as they are

preferred over longer and branched chain alkanes (Giger et al., 1980). The long chain alkanes

have continuously settled out of the water column over time to be incorporated into the sediment

(Huang et al., 2004). These long chain alkanes have obtained a substantial cumulative

concentration almost 20 times that of the short chain alkanes and almost five times the long chain

alkane concentration observed in the oxic zone. The 27 and 29 alkanes are especially indicative

of terrestrial inputs from plants. It is important to note that a lot of the alkanes, especially the

long chained alkane, such as 31 alkane, were not measured at depth because they coeluted with

other more abundant compounds such as alkenes and -OH acids.

Hopanoids have been associated with bacterial activity in the water column or sediment,

and of the reprocessing of primary organic matter (Pearson et al., 2007). This was demonstrated

in Sider’s Pond as the total hopanoids concentration increases with depth along with other

bacterial biomarkers and is highest in the sediment.

13

-OH fatty acids are bacterial biomarkers and have been used as an indicator of strong

sulfate reducing activity in saline aquatic habitats (Goossens et al., 1989; Rotani and Volkman,

2005). Other eukaryotic contributors, such as fungi, have been known to synthesize this

compound class but are comparatively minor (Goossens et al., 1989) and not as likely to be

found in this type of aquatic environment. The concentration of -OH acids increased with depth

indicating the increasing presence of SRB with depth in Sider’s Pond.

The presence of stanols indicated metabolic activity was occurring throughout the water

column, although it was consistently less than the concentration of sterols indicating that

organics are well preserved in this environment. When normalized to POC sterols were lowest at

the oxycline where they may have been reduced in a layer of photosynthetic bacteria such as

Chlorobium, a genus of green sulfur bacteria (Goossens et al., 1989). The low concentration of

stanols relative to sterols in the sediment also suggests that limited diagenesis was occurring

allowing for great sterol preservation (Pearson et al., 2007).

In the surface sediment, long-chain alkanes (>20 carbons) and long, even-chained fatty

acids were observed with chain length distributions very typical of terrestrial plant waxes.

Additionally, large amounts of certain compounds in the sediment are indicative of an

accumulation of products from metabolic and redox reactions occurring throughout the water

column. For example, relatively low concentrations of labile lipids and cholesterol from sinking

feces from zooplankton grazing. Furthermore, C32 hopanol, which is a suspected degradation

product of polyhydroxy-hopanoids or may be synthesized by bacteria (Grimalt et al., 1992;

Rotani and Volkman, 2005), was found only in the sediment.

Conclusions

I found evidence of major differences in pelagic community structure that result from the

unusual meromictic nature of Sider’s Pond. The oxic zone was dominated by planktonic

biomarkers, primarily unsaturated PUFAs. The observed relatively low lipid concentration in the

surface waters also indicated the negative effects of changing environmental conditions as winter

approached on the populations that reside there. Cholesterol also indicated that zooplankton

grazing occurred in the oxic zone. While productivity during this time of year was relatively low

in the surface waters, the oxycline was a hotspot for prokaryotic photosynthesis indicated by

both pigment biomarkers and lipids such as phytol. The anoxic zone was characterized by

branched, odd-chained, saturated fatty acids and -OH fatty acids that were indicative of sulfate

reducing bacteria. Hopanoids also indicated the presence of these sulfate reducers in addition to

bacterial transformational processes occurring at depth. The relatively low concentrations of

labile lipids and overwhelming presence of terrestrial biomarkers in the surface sediment

demonstrates that it provides an excellent record of both autochthonous and allochthonous inputs

and of the transformational processes, often resulting from metabolic bioprocessing of organic

compounds and secondary production, occurring throughout the water column. Furthermore, the

high concentration of cholesterol and low concentration of stanols in the sediment indicated that

14

limited diagenesis was occurring and compounds in the sediment can therefore be well-

preserved.

For future studies I suggest the identification of more biomarkers whether by running the

present samples at higher concentrations to get more resolved MS or using separation and

purification techniques to look at smaller component compounds. I would also suggest sampling

at a multiple times during the year to examine the differences between seasons.

References

Aeckersberg, F., Bak, F., & Widdel, F. 1991. Anaerobic oxidation of saturated hydrocarbons to

CO2 by a new type of sulfate-reducing bacterium. Archives of Microbiology, 156(1):5-14.

Ali, M. M., Humrawali, N., & Latif, M. T. 2009. Phytosterols Composition in Surface Sediment

of Kuala. European Journal of Scientific Research, 33(1):187-194.

Canuel, E. A., Cloern, J. E., Ringelberg, D. B., Guckert, J. B., & Rau, G. H. 1995. Molecular and

isotopic tracers used to examine sources of organic matter and its incorporation into the

food webs of San Francisco Bay. Limonology And Oceanography, 40(1):67-81.

Canuel, E. A., & Martens, C. S. 1996. Reactivity of recently deposited organic matter : near the

sediment-water Degradation interface of lipid compounds. Science, 60(10):1793-1806.

Chen, N., Bianchi, T. S., Mckee, B. A., & Bland, J. M. 2001. Historical trends of hypoxia on the

Louisiana shelf : application of pigments as biomarkers. Carbon, 32:543-561.

Christie, W. 2003. Lipid Analysis: Isolation, Separation, Identification and Structural Analysis of

Lipids. PJ Barnes & Associates: The Oily Press, Bridgwater, UK. pp. 3-36.

Colombo, J. (1996). Lipid biogeochemistry in the Laurentian Trough: I—fatty acids, sterols and

aliphatic hydrocarbons in rapidly settling particles. Organic Geochemistry, 25(3-4): 211-

225.

Franks, J., & Stolz, J. F. 2009. Flat laminated microbial mat communities. Earth-Science

Reviews, 96(3):163-172.

Gilboa-Garber, N. (1971). Direct spectrophotometric determination of inorganic sulfide in

biological materials and in other complex mixtures. Analytical Biochemistry, 43(1): 129-

133.

Goossens, H., Düren, R., Leeuw, J. & Schenck, P. 1989. Lipids and their mode of occurrence in

bacteria and sediments-II. Lipids in the sediment of a stratified, freshwater lake. Organic

Geochemistry 14(1):27-41.

15

Grimalt, J. O., de Wit, R., Teixidor, P., & Albaigés, J. 1992. Lipid biogeochemistry of

Phormidium and Microcoleus mats. Organic Geochemistry, 19(4-6):509–530.

Hama, T. 1991. Production and turnover rates of fatty acids in marine particulate matter through

phytoplankton photosynthesis. Marine Chemistry, 33:213-227.

Huang, Y., Shuman, B., Wang, Y. & Webb III, T. 2004. Hydrogen isotope ratios of individual

lipids in lake sediments as novel tracers of climatic and environmental change: a surface

sediment test. Journal of Paleolimnology 31:363-375.

Lorenzen C. J. 1967. Determination of chlorophyll and pheo-pigments: spectrophotometric

equations, Limnol. Oceanogr., 12(2):343–346.

Meyers, P. 2003. Applications of organic geochemistry to paleolimnological reconstructions: a

summary of examples from the Laurentian Great Lakes. Organic Geochemistry, 34(2):261-

289.

Montoya, J. P., Wiebe, P. H., & McCarthy, J. J. 1992. Natural abundance of 15 N in particulate

nitrogen and zooplankton in the Gulf Stream region and Warm-Core Ring 86A. Deep Sea

Research, 39 (Suppl. I):S363–S392.

Pearson, E., Farrimond, P., & Juggins, S. 2007. Lipid geochemistry of lake sediments from semi-

arid Spain: Relationships with source inputs and environmental factors. Organic

Geochemistry, 38(7):1169-1195.

Pimenov, N. V., Lunina, O. N., Prusakova, T. S., Rusanov, I. I., & Ivanov, M. V. 2008.

Biological fractionation of stable carbon isotopes at the aerobic/anaerobic water interface of

meromictic water bodies. Mikrobiologiia, 77(6):839-847.

Roach, K., Tobler, M., & Winemiller, K. 2011. Hydrogen sulfide, bacteria, and fish - a unique,

subterranean food chain. Ecology, 92(11):2056-2062.

Rontani, J., & Volkman, J. 2005. Lipid characterization of coastal hypersaline cyanobacterial

mats from the Camargue (France). Organic Geochemistry, 36(2):251-272.

Schefuβ, E., Versteegh, G., Jansen, J. & Damsté, J. 2004. Lipid biomarkers as major source and

preservation indicators in SE Atlantic surface sediments. Deep-Sea Research I,

51(9):1199-1228.

Shioi, Y., & Sasa, T. 1984. Terminal Steps of Bacteriochlorophyll a Phytol Formation

Photosynthetic Bacteria. Journal of Bacteriology, 158(1):340-343.

Stal, L., Gemerden, H. V., & Krumbein, W. 1985. Structure and develoment of a benthic marine

microbial mat. FEMS Microbiology Letters, 31:111-125.

16

Villinski, J. C., Hayes, J. M., Brassell, S. C., Riggert, V. L., & Dunbar, R. B. 2008. Sedimentary

sterols as biogeochemical indicators in the Southern Ocean. Organic Geochemistry,

39(5):567-588.

17

Figures and Tables

Figure 1. The temperature (ºC), salinity (ppt), and dissolved oxygen (DO) (mg/L) profile of


Figure 2. The light intensity profile of Sider’s Pond, Falmouth, MA measured in

photosynthetically active radiation (PAR) (µE m-2

s-1

).

Figure 3. The pH profile of Sider’s Pond, Falmouth, MA.

Figure 4. The change in sulfate concentration (µM) with depth at Site 1 in Sider’s Pond,

Falmouth, MA.

Figure 5. The change in sulfide concentration (µM) with depth at Site 1 in Sider’s Pond,

Falmouth, MA.

Figure 6. The change in chlorophyll a concentration (ng/L) with depth at Site 1 in Sider’s Pond,

Falmouth, MA.

Figure 7. The change in concentration (mg/L) with depth for the prokaryotic photosynthetic

pigment biomarkers phaeophytin (indicative of purple sulfur bacteria) and bacteriochlorophyll a.

Figure 8. The change in particulate organic carbon (POC) concentration (µg/L) with depth in


Figure 9. The change in particulate organic nitrogen (PON) concentration (µg/L) with depth in


Figure 10. The change in 13

C fractionation with depth in Sider’s Pond, Falmouth, MA.


N fractionation with depth in Sider’s Pond, Falmouth, MA.

Figure 12. The change in total extractable lipids (TEL) concentration (µg/L) with depth in


Figure 13. The change in total extractable lipids (TEL) concentration after normalization to POC

(µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

Figure 14. The change in total fatty acids (TFA) concentration (µg/L) with depth in Sider’s Pond,

Falmouth, MA.

Figure 15. The change in total fatty acids (TFA) concentration after normalization to POC (ng/µg

Organic C) with depth in Sider’s Pond, Falmouth, MA.

Figure 16. The change in the odd:even fatty acid ratio with depth in Sider’s Pond, Falmouth,

MA.

18

Figure 17. The change in fatty acid saturation ratios (saturated and mono- and polyunsaturated)

with depth in Sider’s Pond, Falmouth, MA.

Figure 18. The change in phytol (squares), and cholesterol (Xs) concentrations (ng/L or µg/g dry

weight) with depth in Sider’s Pond, Falmouth, MA.

Figure 19. The change in phytosterols concentration (ng/L or µg/g dry weight) with depth in


Figure 20. The change in phytol (squares) and cholesterol (Xs) concentrations after

normalization to POC (ng/µg Organic C or µg/mg Organic C) with depth in Sider’s Pond,

Falmouth, MA.

Figure 21. The change in phytosterol concentrations after normalization to POC (ng/µg Organic

C or µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

Figure 22. The change in the concentrations (ng/L or µg/g dry weight) of the phytosterols;

cholesta-5,22E-dien-3-ol (diamonds), brassicasterol (squares), brassicastanol (triangles), -

sitosterol (Xs), and campesterol (circles) with depth in Sider’s Pond, Falmouth, MA.

Figure 23. The change in the concentrations (ng/L or µg/g dry weight) of the phytosterol,

stigmasterol with depth in Sider’s Pond, Falmouth, MA.

Figure 24. The change in alkane concentration (ng/L or µg/g dry weight) with depth in Sider’s

Pond, Falmouth, MA.

Figure 25. The change in alkane concentration after normalization to POC (ng/µg Organic C or

µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

Figure 26. The change in total hopanoids concentration (ng/L or µg/g dry weight) with depth in


Figure 27. The change in total -OH fatty acids concentration (ng/L or µg/g dry weight) with

depth in Sider’s Pond, Falmouth, MA.

Figure 28. The change in total hopanoids concentration after normalization to POC (ng/µg

Organic C or µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

Figure 29. The change in total -OH fatty acids concentration after normalization to POC (ng/µg


Figure 30. The change in total sterols and stanols concentration (ng/L or µg/g dry weight) with


19

Figure 31. The change in total sterols and stanols concentration after normalization to POC

(ng/µg Organic C or µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

Table 1. The lipid biomarkers commonly associated to certain groups of organisms that are

characteristic of an ecosystem like Sider’s Pond, Falmouth, MA.

Table 2. The change in saturation and odd:even ratios of fatty acids with depth in Sider’s Pond,

Falmouth, MA.

Table 3. The concentrations of all observed alkanes throughout the water column.

Table 4. The variation in alkane chain length [short chain (<20) versus long chain (>20)]

throughout the water column.

20

Figure 1. The temperature (ºC), salinity (ppt), and dissolved oxygen (DO) (mg/L) profile of


0

1

2

3

4

5

6

7

8

9

10

11

12

13

0 5 10 15 20 25 D

epth

(m

)

Temperature (°C)

Salinity (ppt)

DO (mg/L)

21

Figure 2. The light intensity profile of Sider’s Pond, Falmouth, MA measured in

photosynthetically active radiation (PAR) (µE m-2

s-1

).

0

1

2

3

4

5

6

7

8

9

10

11

12

13

0 200 400 600 800 1000 1200 1400 D

epth

(m

) PAR (µE/m2/s)

22

Figure 3. The pH profile of Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

6.6 6.8 7.0 7.2 7.4 7.6

Dep

th (

m)

pH

23

Figure 4. The change in sulfate concentration (µM) with depth at Site 1 in Sider’s Pond,

Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

0 2000 4000 6000 8000 10000 12000 D

epth

(m

) Sulfate concentration (µM)

24

Figure 5. The change in sulfide concentration (µM) with depth at Site 1 in Sider’s Pond,

Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

0 1 1 2 2 3 3 4 4 5 D

epth

(m

) Sulfide concentration (µM)

25

Figure 6. The change in chlorophyll a concentration (ng/L) with depth at Site 1 in Sider’s

Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

0 5 10 15 20

De

pth

(m

) Chlorophyll a concentration (µg/L)

26

Figure 7. The change in concentration (mg/L) with depth for the prokaryotic

photosynthetic pigment biomarkers phaeophytin (indicative of purple sulfur bacteria) and

bacteriochlorophyll a.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 5 10 15 20 25

Dep

th (

m)

Concentration (mg/L)

Phaeophytin (Purple Sulfur Bacteria)

Chl-GSB (Green Sulfur Bacteria)

Bacteriochlorophyll

a

27

Figure 8. The change in particulate organic carbon (POC) concentration (µg/L) with depth

in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 500 1000 1500 2000 2500 3000

Dep

th (

m)

µg Organic C / L

Sediment Carbon = 139.1 ± 1.6 mg/ g dry wt

28

Figure 9. The change in particulate organic nitrogen (PON) concentration (µg/L) with


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 100 200 300 400 500 600

Dep

th (

m)

µg Organic N / L

Sediment Nitrogen = 14.0 ± 0.2 mg/g dry wt

29


C fractionation with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

-37 -35 -33 -31 -29 -27 -25

Dep

th (m

) 13C (‰)

Suspended Particles

Sediment

30

Figure 11. The

N fractionation with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 2 4 6 8 10 12 D

ep

th (

m)

15N (‰)

Suspended Particles

Sediment

31

Figure 12. The change in total extractable lipids (TEL) concentration (µg/L) with depth in


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 50 100 150 200 250 300

Dep

th (

m)

TEL (µg/L)

Sediment = 7175.73 ± 511.57 mg/g dry wgt

32

Figure 13. The change in total extractable lipids (TEL) concentration after normalization

to POC (µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0.00 0.05 0.10 0.15

Dep

th (

m)

TEL (µg TEL/µg Organic C)

Sediment = 51.58 ± 4.26 µg/mg dry wgt

33

Figure 14. The change in total fatty acids (TFA) concentration (µg/L) with depth in Sider’s

Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 20 40 60 80 100 120

Dep

th (

m)

Concentration (µg/L)

TFA

Sedimment = 2443.27 ± 354.45 µg/g dry weight

34

Figure 15. The change in total fatty acids (TFA) concentration after normalization to POC

(ng/µg Organic C) with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 10 20 30 40 50 60 70 80 90

Concentration (ng/µg Organic C)

TFA

Sediment = 17.56 ± 1.6 µg/mg Organic C

35

Figure 16. The change in the odd:even fatty acid ratio with depth in Sider’s Pond,

Falmouth, MA.

0% 20% 40% 60% 80% 100%

2

5.5

10

12

Sediment

Percent Distribution D

ep

th (

m)

Odd Chain

Even Chain

36

Figure 17. The change in fatty acid saturation ratios (saturated and mono- and

polyunsaturated) with depth in Sider’s Pond, Falmouth, MA.

0% 20% 40% 60% 80%

2.0

5.5

10.0

12.0

Sediment (12.5m)

Percent of Total Fatty Acids D

epth

(m

)

Polyunsaturated

Monounsaturated

Saturated

37

Figure 18. The change in phytol (squares), and cholesterol (Xs) concentrations (ng/L or

µg/g dry weight) with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 100 200 300 400 500 600 700

Dep

th (

m)

Concentration (ng/L)

Phytol

Cholesterol

Sediment (µg/g dw): Phytol = 39.49 ± 10.27 Cholesterol = 37.04 ± 3.89

38

Figure 19. The change in phytosterols concentration (ng/L or µg/g dry weight) with depth

in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 2000 4000 6000 8000

Dep

th (

m)

Phytosterol concentration (ng/L)

Sediment: = 225.92 ± 10.87 µg/g dw

39

Figure 20. The change in phytol (squares) and cholesterol (Xs) concentrations after

normalization to POC (ng/µg Organic C or µg/mg Organic C) with depth in Sider’s Pond,

Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0.0 0.1 0.2 0.3 0.4 0.5 0.6

Dep

th (

m)


Phytol

Cholesterol

Sediment (µg/mg Org C) : Phytol = 0.284 ± 0.077 Cholesterol = 0.266 ± .031

40

Figure 21. The change in phytosterol concentrations after normalization to POC (ng/µg


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 1 2 3 4 5 6

Dep

th (

m)

Phytosterol concentration (ng/µg Organic C)

Sediment = 2.09 ± 0.04 µg/mg Org C

41

Figure 22. The change in the concentrations (ng/L or µg/g dry weight) of the phytosterols;

cholesta-5,22E-dien-3-ol (diamonds), brassicasterol (squares), brassicastanol (triangles),

-sitosterol (Xs), and campesterol (circles) with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 1000 2000 3000 4000 5000

Dep

th (

m)

Phytosterol concentration (ng/L)

Cholesta-5,22E-dien-3ß-ol

Brassicasterol

Brassicastanol

ß-Sitosterol

Campesterol

Sediment (µg/g dw): Cholesta-5,22E-dien-3-ol = 60.89 ± 5.24

-sitosterol = 64.53 ± 0.44 Brassicasterol = 0.00

Campesterol = 97.38 ± 1.62 Brassicastanol = 7.47

42

Figure 23. The change in the concentrations (ng/L or µg/g dry weight) of the phytosterol,

stigmasterol with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 1 2 3 4 5

Dep

th (

m)

Stigmasterol concentration (ng/L)

Sediment = 60.18 ± 4.01 µg/g dw

43

Figure 24. The change in alkane concentration (ng/L or µg/g dry weight) with depth in


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 100 200 300 400

Dep

th (

m)

Alkane concentration (ng/L)

Sediment = 165.05 ± 22.53 µg/g dw

44

Figure 25. The change in alkane concentration after normalization to POC (ng/µg Organic

C or µg/mg Organic C) with depth in Sider’s Pond, Falmouth, MA.

0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0.0 0.1 0.2 0.3 0.4

Dep

th (

m)

Alkane concentration (ng/µg Organic C)


45

Figure 26. The change in total hopanoids concentration (ng/L or µg/g dry weight) with


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 200 400 600 800 1000

Dep

th (

m)

Hopanoid concentration (ng/L)

Sediment = 95.21 ± 15.58 µg/g dry weight

46

Figure 27. T -OH fatty acids concentration (ng/L or µg/g dry weight)


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 1000 2000 3000 4000 5000 6000

Dep

th (

m)

-OH Acid concentration (ng/L)

Sediment = 119.23 ± 21.53 µg/g dry weight

47

Figure 28. The change in total hopanoids concentration after normalization to POC (ng/µg


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0.0 0.1 0.2 0.3 0.4 0.5 0.6

Dep

th (

m)

Hopanoid concentration (ng/µg Organic C)


48

Figure 29 -OH fatty acids concentration after normalization to POC


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 1 2 3 4 5

Dep

th (

m)

-OH concentration (ng/µg Organic C)


49

Figure 30. The change in total sterols and stanols concentration (µg/L or µg/g dry weight)


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 1 2 3 4 5 6 7 8

Dep

th (

m)

Concentration (µg/L)

Other Sterols

Stanols

Sediment (µg/g dw) = Sterols = 642.03 ± 52.79 Stanols = 48.60 ± 2.56

50

Figure 31. The change in total sterols and stanols concentration after normalization to POC


0

1

2

3

4

5

6

7

8

9

10

11

12

13

14

0 2 4 6 8 10 12

Dep

th (

m)


Other Sterols

Stanols

Sediment (µg/mg Organic C) = Sterols = 4.61 ± 0.11 Stanols = 0.35± 0.03

51

Table 1. The lipid biomarkers commonly associated to certain groups of organisms that are

characteristic of an ecosystem like Sider’s Pond, Falmouth, MA.

Compounds Eukaryotic

phytoplankton

Zooplankton Phototrophic

bacteria

Sulfate reducing

bacteria

Odd:Even Fatty

Acid Ratio

↓ ↓ ↑ ↑

Polyunsaturated

Fatty Acids

(PUFAs)

↑ ↑ ↓ ↓

Phytosterols ↑ ↓ - -

Phytol ↑ - ↑ -

Cholesterol - ↑ - -

Hopanoids and

-OH Acids

- - ↑ ↑

52

Table 2. The change in saturation and odd:even ratios of fatty acids with depth in Sider’s

Pond, Falmouth, MA.

EVEN ODD

Depth

(m)

Poly-

unsaturated

(%)

Mono-

unsaturate

d (%)

Saturated

(%)

Poly-

unsaturated

(%)

Mono-

unsaturate

d (%)

Saturated

(%)

2 33.2 31.6 31.2 0.0 0.0 3.9

5.5 21.2 17.0 45.9 0.9 0.2 14.7

10 5.2 20.8 42.9 1.8 0.6 28.7

12 6.5 21.5 39.4 3.9 1.3 27.4

Sediment 6.6 22.6 58.6 3.4 0.6 7.5

53

Table 3. The concentrations (ng/L) of all observed alkanes throughout the water column.

Depth (m) C16 C17 C21 C23 C24 C25 C26 C27 C28 C29 C38

2 0.0 81.2 0.0 0.0 0.0 0.0 0.0 34.6 0.0 0.0 10.4

5.5 0.0 136.5 0.0 0.0 0.0 0.0 0.0 49.6 0.0 0.0 34.5

10 67.5 168.2 0.0 0.0 0.0 0.0 42.1 0.0 0.0 0.0 10.3

12 93.7 243.8 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 10.0

Sediment 0.0 7.0 13.6 9.2 1.7 17.6 0.0 37.6 33.5 43.0 1.9

54

Table 4. The variation in alkane chain length [short chain (<20) versus long chain (>20)]

throughout the water column.

Depth (m) < 20 C (ng/L) >20 C (ng/L)

2 81.2 26.8

5.5 136.5 84.1

10 235.7 52.5

12 337.5 10.0

Sediment (µg/g dw) 7.0 120.5

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