A week before, arrange use of fluormeter, DOC machine and other equipment with other users. Check if supplies or chemicals need to ordered. The day before prepare be sure solutions are prepared.

Before leaving the lab, set the incubators to the in situ temperature is you know this already. This will allow the incubator to be at the correct temperature by the time you need them. If you do not know the in-situ temperature, be sure to ask the LTER base crew if you will see them on the lake. If they will not be on the lake, borrow a temperature probe and take the temperature at the middle of the epilimnion.

At the deep hole station, deploy the O2 sonde for the lake metabolism measurements before sampling.

Integrated samples of the epilimnion of each study lake are collected at the same day as LTER base crew sampling, from the deep hole station. The epilimnion is defined as the surface water extending down to the depth at which temperature changes more than one degree centigrade over one meter. For Crystal Bog the sampling depth is the entire water column since it does not stratify. The tube is first rinsed with lake water by lowering one end of the tube to the bottom of the epilimnion with both ends open. Samples can now be collected. The first sample is used to rinse the collection containers and is then discarded.

To collect samples, one end of a tube is lowered to the bottom of the epilimnion. The top end is then closed and the lower end retrieved by the attached rope. At Trout Lake the PVC integrator tube is used. The sample is poured into a sampling container through a 70 µm mesh sieve to collect the large zooplankton. Separate sample hauls are put into separate containers, for a total of three sample replicates. The same 70 µm mesh sieve is used for all samples without rinsing between samples in order to collect the large zooplankton. When all the samples have been collected the sieve is covered with foil or put in a bag and stored on ice in the cooler.

The procedures for processing of samples is detailed in the following sections of this methods book. The general order of dealing with samples is as follows. Back at the lab, aliquots are taken for bacterial counts, heterotrophic nanoflagellates, algae and protists. Zooplankton are removed from the sieve and collected. Gluteraldehyde or other appropriate poison is added, per each method.

Next 250 mL of sample filtered through a 0.2 mm filter for dissolved chemistry parameters (DOC, DAC, protein, spectra). Then 125 ml of sample is filtered through a 1 mm filter for the <1 µm bacterial count, active bacterial counts, ETS, APA and bacterial production parameters. Gluteraldehyde is added to the bacterial count aliquots.

The remaining material is used for various incubations. The remaining sample should be stored in an incubator set to in situ temperatures. Start the longest incubations first.

Incubations are completed the same day except for planktonic. The final O₂ measurement for planktonic respiration is taken after 24 or 48 hours. The O₂ sonde measuring lake metabolism must be retrieved from the field after a few days.

Once incubations are done then SUVA/Spectral Analysis is measured at ambient pH and after pH has been adjusted to 7.0.

At Trout Lake, the preserved BN, HNF, Zooplankton, Protist-Algae samples, and the <0.2 mm water prepared for DOC/Protein/Spectral Analysis is stored in a frig and shipped on ice to Madison at the earliest opportunity. The DNA samples are deep frozen and also shipped to Madison at the earliest opportunity. FISH slides are fixed and prepared up to the ethanol rinse, then stored in a slide box at room temperature until shipped to Madison. Bacterial production samples are incubated at Trout Lake. After stopping with 50% TCA, the samples in microcentrifuge tubes are stored in a frig until transferred to Madison for final processing. Planktonic respiration is completely conducted at Trout Lake. Alkaline phosphatase (APA) and Lysozyme activity can not currently be conducted at Trout Lake.

SUMMARY OF ANALYSES

ALL samples field filtered through 70 µm mesh

Unfiltered lake water

DNA (Eric)
FISH	6 slides
BN	3 replicates
HNF	3
Protists & algae (Jim)	1
BP	4 + 2 poisons
BOD (Winklers)	7 initial, 7 final
lysozyme	3
Phosphatase (APA)	3

<1 µm lake water

BN	3
BP	4 + 2 poisons
Respiration	7 initial, 7 final
APA	3

<0.2 µm lake water

Protein	3
DOC	3
Spectral Analysis	6 (three at ambient pH and three at pH=7)
APA	3
ADOC	3

Additional items

Zooplankton	1
Lake metabolism (O2 sonde) deployed five days

MICROBIAL ABUNDANCE, IDENTIFICATION AND BIOMASS

FLUORESCENT IN SITU HYBRIDIZATION (FISH)

SLIDE PREPARATION (Day of sampling)

Solutions

use Milli-Q fresh from the tap for all solutions

EtOH-KOH slide wash: 95% ethanol, 10 % KOH, add more water if needed to get in solution.

3X PBS: NaCl (Fisher BP358-1), Na phosphates (Fisher BP332-500, BP329-500)

Geletin solution: 0.075% gelatin (0.15g) (Fisher G-9391), 0.01% chromium potassium sulfate dodecahydrate (0.02g) (Sigma L-5926 ACS)

4% paraformaldehyde solution (prepare daily, stable 48 hours, Sigma P-6148): 3X PBS, 10 N NaOH, 1 N Hcl

lysozyme solution: 12.5 mg lysozyme in 25 mL solution (Sigma L-7651)

EtOH washes: 50, 80, 96 %

Fuge method also requires: 1X PBS

Supplies

Small and large pipet tips

pipettors capable of 5 to 1000 µL volumes

solution bottles

slides

slide holders

slide stain container: EtOH washes, slide prep

weigh boats

filtration manifold

0.2 µm CTE filter, 25 mm diameter

tweezers

vacuum dessicator

37 C oven

pH meter

petri dishes for fixation

Milli-Q squirt bottle

Fuge method also requires: centrifuge, vortexer, 2 mL centrifuge vials, centrifuge vial holders

Recipes

Gelatin

Prepare 0.15 g gelatin (0.075%)and 0.02 g chromium potassium sulfate dodecahydrate (0.01 %) in 200 g of nanopure water. Heat to 70C to dissolve gelatin. Dip slides in heated gelatin and dry.

Fixation Solution (4% Paraformaldehyde)

2 g paraformaldehyde

20 µl 10 N NaOH

16.5 mL nanopure water

Heat above to 60 C until dissolves

Add 33 ml of 3X PBS and cool to 4 C

Adjust pH to 7.2 with 1 N HCl

Filter through 0.2 µm syringe filter

Keep refrigerated. Stable for 48 hours.

3 X Phosphate Buffer Solution (PBS)

Prepare 1 M solutions of Na₂HPO₄ and NaH₂PO₄.

For 100 mM, in 1000 mL add 68.4 mL 1M Na₂HPO₄ and 31.6 mL 1M NaH₂PO₄.

390 mM NaCl

30 mM sodium phosphate

Adjust pH to 7.2

Lysozyme

Prepare 0.5 mg ml-1 by adding 12.5 mg in 25 mL Nanopure water.

Procedure

WASH slides in EtOH-KOH for one hour. Rinse well with Nanopure squirt bottle and air dry. Coat slides by dipping in geletin solution heated to 70 C (heating setting is about 2.3). Let air dry by leaning one end on weigh boat, upside down to minimize dust settling on surface. You can prepare a bunch of slides ahead of time. Store in a separate slide holder marked to indicate these are coated but unused slides.

PREPARE a total of six slides per lake on each collection date. That is a total of 48 wells.

Filtered Sample Prep

FILTER sample onto a 0.2 µm PCTE filter, 47 mm diameter. Aliquot 5 the µL Nanopure water in eight wells on slide to help the filter stick. Place filter onto coated slide covering eight wells. Do not use the two wells on the end. Press filter down gently but firmly with thumb. Dry 30-120 minutes, in vacuum dissicator if possible. While drying, prepare fixation (4% paraformaldehyde) and lysozyme solutions.

LIFT filter off and discard. Add 20 µL of 4% paraformaldehyde solution to each well. Incubate for 30 minutes at room temperature, covered in petri dish to prevent dust and drying.

RINSE off 4% paraformaldehyde solution thoroughly with Nanopure water squirt bottle. Continue with final slide preparation procedures.

Aqueous Sample Prep (not used anymore-appears too selective)

CENTRIFUGE 1.8 mL of lake water at 14,000 RPM for 4 minutes. Remove top 1.5 mL.

ADD 900 µL (three volumes) of 4% paraformaldehyde solution. Vortex for 10 seconds. Incubate 30 minutes at room temperature.

CENTRIFUGE at 14,000 RPM for 4 minutes. Decant paraformaldehyde solution. Add 1 mL of 1X PBS for a wash and vortex for 10 seconds.

CENTRIFUGE at 14,000 RPM for 4 minutes. Decant PBS solution. Add 1 mL of 1X PBS and vortex for 10 seconds.

ALIQUOT 5 mL of sample to each well. Do not use wells on end. Let air dry. Continue with final slide preparation procedures.

Final Slide Preparation

TREAT one slide with lysozyme by adding 30 µl of lysozyme solution. Incubate 30 minutes at 37 C. Rinse off solution thoroughly with Milli-Q water.

DEHYDRATE all slides by immersing in 50, 80 and 96 % ethanol for 2 minutes each, then air dry and store in slide holder. All slides should be marked legibily with lake abbreviation, collection date, “lys” if lysozyme treated, and volume of lake water filtered or centrifuged if different than procedure above. Slides should also be individually numbered (i.e. 1 to 6) to differentiate between slides created on same date. Slides prepared at Trout Lake are stored in a slide holder at this point and shipped to Madison at the next opportunity.

PROBE HYBRIDIZATION

Supplies

50 mL centrifuge tubes, two per slide

46-48 C incubator

1 mL pipetor

20 uL pipetor

paper towels

Solutions

0.9 M NaCL

5 M NaCl

1 M Tris-HCl

10 % SDS

probes, prepared at 100 ug/ mL in TE, pH 8.0

DAPI, 1 ug/ mL

Antifadent (eg. Citifluor, DABCO, etc.)

Recipes

Procedure

DAY BEFORE determine slide probe locations and conditions. Calculate hybridization and wash buffer conditions,

START oven at 46 C. Moisten paper towels with 0.9 M NaCl and place in 50 mL polypropylene centrifuge tubes, one tbe for each set of hybrization conditions.

PREPARE hybridization buffer.

CONDUCT rest of method in low light conditions.

ADD 16 ul of hybridization buffer and 0.5 uL of probe to each well. Place slide in preincubated moisture chamber and incubate for 1.5 hours at 46 C.

PREPARE washing buffers and fill 50 mL centrifuge tubes to 35 mL mark. Preincubate at 46 C.

REMOVE slides from incubator and change incubator temperature to 48 C.

RINSE slides with 1 ml of wash buffer twice. Immerse slides in wash buffer and incubate for 20 min at 48 C.

RINSE slides with 1 mL of Nanopure water, then air dry. You can store slides at room temperature at this point.

COUNTERSTAIN with DAPI by adding 40 u: of 1 ug/ mL DAPI solution to each well. Let sit 5 min then rinse briefly with Nanopure water.

ADD 5 uL of antifadent (eg. Citifluor, DABCO, etc.) to each well, then cover with a large coverslip. Carefully seal edges with nail polish unless viewing immediately.

EXAMINE slide under proper fluorescence microscope conditions. Using paper, count each cell first under probe excitation, then as DAPI cells, by noting location with a pencil mark. Save an electronic copy of each image.

BACTERIAL COUNTS (BN)

Supplies

scintillation vials

gluteraldehyde, add 0.8 ml to 20 ml of sample

25 mm glass filtration set: top, base, clamp, vacuum flask

vacuum source, set to 10 cm Hg

0.2 µm polycarbonate track-etched black filter, 25 mm

tweezers, flat tip

glass slides

cover slips

slide holder

DAPI, 10 ug ml-1

syringe, with 0.2 µm filter tip, for DAPI

syringe, with 0.2 µm filter tip, for Nanopure water

Sample preparation

Label vial cap with lake abbreviation, date, and "BN" and the replicate number. Aliquot 20 ml of lake water sample into new scintillation vial. Add add 0.8 ml gluteraldehyde to 20 ml of sample. Store at 4^o C.

Filtration

TURN down lights in room as much as possible. DAPI is somewhat light sensitive. Pre-rinse filter set-up with 0.2 µm filtered Nanopure water.

PUT filter on filtration base with vacuum on, and clamp on filtration top. The vaccum helps filter sit flat. Turn off vacuum.

ALIQUOT	appropriate sample volume
eutrophic lakes (ME)	use 0.5 ml
humic lakes (CB, MA)	use 1 ml
oligotrophic lakes (CR)	use 2 ml

ADD DAPI, 7 drops per ml of sample.

COVER with foil. Let sit while you prepare slide.

PREPARE slide. Using black Sharpie, label with lake name, date, BN and replicate number (1-3). Place drop of immersion oil in center and spread as evenly as possible in a square 3 cm area.

TURN on vacuum. When less than 1 ml of water is left to filter, rinse filtration top with Nanopure water, being sure to get all walls. This will break up the surface tension and help cells lay evenly on filter and remove excess DAPI.

REMOVE filter while vacuum is still on Once all the water has been filtered, and then wave the filter in the air to a count of twenty to completely dry it.

PLACE filter on prepared slide. Place a drop of oil in center of filter and cover with coverslip. Let oil distribute evenly by gravity while you prepare next filter on filter unit (step 2).

COVER slides with paper towel and gently press down to remove excess oil. Store finished slides in slide holder in freezer.

CONTINUE with other slides.

CHECK finished slides briefly to be sure DAPI fluorescence is strong enough and cells are evenly distributed. Samples should only be tossed when slides have been counted.

BACTERIAL ABUNDANCE - counting

Counting

BRING slides, Nikon immersion oil, keys and radio to Center for Limnology Lab.

STORE slides in freezer in lab. Remove first two slides to thaw while you get the microscope ready. Always let the next slide thaw as you are counting the current slide.

TURN on mercury lamp by flipping switch on box labelled CHIU Technical Mercury 100 W. Sign in the notebook your name and the time you turned on the lamp. Note that the lamp must not be turned back off for a least thirty minutes to prevent damage.

PUT orange visor above oculars to protect from UV light damage to eyes. Turn off room lights if no one else minds. Check that both photo knobs are pushed in on right side of microscope.

BRING the lamp alignment tool objective into position. If white, diffuse light is not visible in tool check that shutter is open. The shutter is located between the lamp and the microscope and slides out to the left. Locate the largest knob on the top of the lamp housing located on the right side of the lamp box. This knob moves the lamp from side to side in the tool. The knob on the side of the lamp housing moves the lamp up and down in the tool. The two smaller, off-center knobs on the lamp housing hold the lamp in and should not be adjusted. Center the lamp in the tool. Using the large knob located in front of the lamp housing behing the microscope to bring the lamp coils into focus as best you can.

POSITION the slide upside-down on microscope stage. The slide will just fit in the recessed opening lip. Turn on the visible light lamp by the switch on the lower left front of the microscope. Using the 40X objective find the focal plane by focusing on the black filter particles. Turn off visible light lamp and should see the blue-white constellation of bacterial cells and other organisms.

TURN 100 X objective halfway into position. Drop a dab of Nikon immersion oil directly onto objective and bring into position. Using the stage controls, move the stage around so that the oil is spread around from the tip of the 100X objective.

DETERMINE about how many cells are in a Wipple grid. Choose the subset of the grid to count that holds about 30 cells (range 20-60). The different seubsets are the whole grid, 3/4 grid, 1/2 grid and 1/4 grid. If there are too few cells or two many then redo slide.

COUNT all the cells in each Wipple grid subset in ten places randomly choosen along two transects across slide, five on each transect. Use a total of two transects, which will cross filter perpendicular to each other. Start and end each transect near the edge of the filtration area, i.e. where there are no longer cells. In notebook, note the sample name, number of cells in each field, the total number of cells in 10 fields, the size of the Wipple grid counted (1,3/4,1/2,1/4), and photo number if photo was taken.

WHEN done, store slides in freezer and take out another slide to thaw. When done for the day, put all slides in slide box and store in freezer. Turn off mercury lamp and note time in lamp notebook. Clean excess oil off 100 X objective carefully using lens paper in lamp notebook. Uses clockwise motion as you wipe around the lenses.

HETEROTROPHIC NANOFLAGELLATES (HNF)

Heterotrophic nanoflagellates samples and slides are prepared the same way as the bacterial count samples and slides. The only differences are the use of a 0.8 µm filter, a larger volume of 10 ml, and a stronger DAPI concentration of 100 µl of 0.1 mg ml-1 DAPI for every millileter of sample. Sample can be drawn down to approximately two ml before adding 200 µl of 0.1 mg ml-1 DAPI. Make 0.1 mg DAPI mL-1 by diluting 2 ml of 1mg mL-1 DAPI in 18 mL of Nanopure water. Keep in dark.

To count, use the same method as for bacteria, but count using the 40 X, non-oil objective. Again, count ten fields of about 30 cells each. Note anything interesting you see about organisms and what their cells are associated with. Interesting things to note may include detritus or colonies.

ZOOPLANKTON SIZE

Zooplankton captured on the 70 µm sieve are rinsed off using Nanopure water into a scintillation vial. Scintillation vials are prelabeled with lake abbreviation, date and "Zoops". Lake water is added to bring sample up to approximately 20 ml, at the shoulder of the vial. 0.8 ml of 50 % gluteraldehyde is added to preserve sample. Sample is stored in cold room. All zooplankton are identified and sized.

ALGAE AND PROTISTS

250 ml of lake sample is aliquoted into a polyethelyene bottle. Gluteraldehyde is added to a final concentration of 1 %. Sample is stored in cold room till it can be delivered to Jim. Bottle should be labeled with lake abbraviation and date.

DNA COLLECTION

Supplies

2 bottles (glass or plastic)

10 um nylon mesh (4 square inches)

cylinder

large funnel (funnel must be small enough to fit inside bottle cap and large enough to encompass the mesh/cylinder)

sterile DI water

cooler for the samples

sampling device (usually 2 reps required to obtain a full 2 liters of water)

1 liter side arm flask

vacuum pump.

bottle top filter set up.

screw cap

sterilized tweezers

0.2 um SUPOR ployethersulfone filters.

screw top 250ml collector.

cryo-vials and label with volume, date and lake name.

–80 degree cold storage

Preparations

Sterilize 2 bottles (glass or plastic)

Cut and bag 10 um nylon mesh (4 square inches)

Place mesh into cylinder to secure

Acid wash large funnel and rinse with sterile DI water

(funnel must be small enough to fit inside bottle cap and large enough to encompass the mesh/cylinder)

Get a cooler for the samples

Field Sampling

Collect water with sampling device (usually 2 reps required to obtain a full 2 liters of water)

Place hose of sampling device so the water flows through the cylinder/mesh-funnel and into the bottle. Go slowly

Collect one liter clamp off hose, cap first bottle and place into cooler. Repeat with second bottle.

Sample Processing

Remove bottles of water and place into refrigerator

Set up bacterial filtering apparatus

Attach side arm of 1 liter side arm flask to a vacuum pump.

A top the side arm flask place bottle top filter set up.

First place screw cap on and using a sterilized tweezers place 0.2 um SUPOR ployethersulfone filters.

Attach screw top 250ml collector.

Apply water into collector and turn on vacuum pump

Run 500ml of water through each filter-depending of lake type (crystal bog 250mL)

Turn off vacuum pump and using flame sterilized tweezers remove and fold bacterial filter

Place filter into cryo-vials and label with volume, date and lake name.

Place into –80 degree cold storage

Repeat with next batch of water and new filter

MICROBIAL ACTIVITY

PLANKTONIC RESPIRATION (BOD)

Supplies

14 BOD bottles of same size and same stopper size, weighed semi-annually to 0.01 g

peristaltic pump with tubing

Cooler with ice and sample

MnSO4 (Fisher/Lab Chem LC16570-4)

NaOH-NaN3-NaI (Fisher SA435-1)

H2SO4, concentrated (Fisher A484-212)

Na2S2O3 (Fisher SS370-1)

starch indicator

10 ml buret with stopcock and stand

10 ml pipet with bulb for filling buret

pasteur pipet for starch indicator

Erylenmeyer flask, 500 ml

Procedure

FILL 14 BOD bottles with sample using a peristaltic pump. Use tubing to add lake sample at the bottom surface of the BOD bottle. Let three volumes pass through bottle. Remove tubing slowly being careful to keep bottle filled to rim and not introduce bubbles. Insert stopper being careful not to leave bubbles. This requires some practice. I use a quick twisting motion while the bottle is held at a slight angle.

PLACE seven BOD bottles in incubator set to in situ temperature and darkness. These will be your final time point samples. If lake is oligotrophic or in situ temperature is < 15^oC, incubate 48 hours. Otherwise incubate only 24 hours.

TURN on spectrometer to warm up lamp. Put on gloves.

USING a micropipet add 1 ml of MnSO4 just above the surface, then 1 ml of NaOH-NaI-NaN3 just below the surface. Insert cap again being careful not to introduce air. Hold lid with thumb and shake vigorously. Do all seven bottles. Note time in notebook. Wait till precipitate has settled down to the bottom third of bottle. Shake bottle to resuspend the precipitate a second time. Wait till precipitate has settled down to the bottom third of bottle again before adding acid.

ADD 1 ml concentrated H2SO4 using a micropipet. Insert cap and shake as above. Rinse micropipet outside and chamber thoroughly after use. If you must pause before analysing, store bottle at temperature slightly lower then the sample temperature.

ANALYZE seven initial and seven final bottles using the spectroscopic method. Also analyze four intial and four final bottles using the titration method. To do both analyses, remove 10 mL for spectroscopic analysis before titrating the rest.

Spectroscopic Analysis

ZERO spectrometer with Nanopure water. Measure absorbance of sample at 430 nm wavelength in a 10 mm glass cuvette. Use Nanopure water in the reference cell if applicable. Rinse cuvette thoroughly with Nanopure water between samples. Measure each sample twice.

CALCULATE dissolved oxygen from the equation

DO (mg L-1) = 0.0081 * absorbance – 0.410

[accurate for DO between 4 and 12 mg L-1, from Roland et al., Limnol Oceanogr., 1999, 44:1148-1154]

DON'T FORGET you have to do seven more bottles at 24 or 48 hours from inital time point.

Titration Analysis

ONLY titrate four bottles each from the initial and final sample pools. Titrate with sodium thiosulfate (Na2S2O3) and strach indicator. This is the delicate part of the procedure, so take your time. The bottles should be allowed to come to room temperature before titrating. Fill buret with about 10 ml sodium thiosulfate and note volume in notebook. Pour sample into Erylenmeyer flask. Slowly add thiosulfate until solution changes from a rusty orange to a straw yellow.

ADD at least 10 drops of starch indicator to turn solution blue. Continue titrating but only drop by drop. Once solution is a deep blue you are within 1 ml of end. Add thiosulfate drop-by-drop till all blue color disappears. Note final volume from buret. Empty flask and rinse with distilled or reverse-osmosis (RO) water three times before next sample.

BOG water is more difficult to titrate because it always has a background yellow color that makes endpoints harder to see. Do your best and take your time. I find that the replicate bottles are always within a milliliter of each other, so once the first one is done you have a general idea how much titratant for other three bottles.

DON'T FORGET you have to do four more bottles at 24 or 48 hours from inital time point.

BACTERIAL PRODUCTION (³H-LEUCINE)

Supplies

50% trichloroacetic acid (TCA, Fisher SA433-500))

3H-leucine

Leucine, non-radioactive

2 mL microcentrifuge tubes with o-rings

microcentrifuge tube holders

radioactive labelling tape

5% TCA

Recipes

3H-Leucine

5-29: Conc. Leu: about 32 mg leucine in 100mL, removed 60 mL and diluted remaining 40 mL with 60 mL Nanopure.

PREPARE 3H-leucine working solution. First prepare concentrated leucine solution, 1.28 mM leucine, by dissolving 167.897 mg leu in 1 liter. For the 3H-leucine working solution (320 nM), prepare 20 mL of Nanopure water in a scintillation vial. Remove 55 µL. Add 5 µL of concentrated leucine stock and 50 µL of 3H-leucine stock (1 mCi mL-1). Concentrated and working solutions are good for two sampling periods (three weeks).

Equipment

Incubator

Pipettor capable of 100 to 1,500 µL volumes