Research Recipes

A repository for my research protocols

    Granted, some of these recipes are more for me to look up to remind me of protocols, but if you can make use of them, or better yet suggest some improvements, please let me know!
In this section:

Cephalopod anaesthetics
  1. Measure out 3L of holding seawater into a 5 gallon plastic pail
  2. Place 100-300 gm octopus into bucket
  3. Slowly dribble 3% of 95% Ethanol (i.e. 90 mL) down the wall of the bucket.
  4. Wait for appropriate octopus anaesthetization
Cephalopod ringers for E-physiology

(Ingredient)(in mmol/L)(in g/L)
NaCl47027.47g
KCl100.75g
CaCl2.6H2O602.19g
MgCl2.6H2O5010.17g
Glucose203.60g
HEPES102.38g

Slowly adjust to pH 7.8 using 2.0 N NaOH.

Cephalopod buccal mass paraffin sectioning
From Kier WM. 1992. Hydrostatic skeletons and muscular hydrostats, Ch. 9. In: Biewener AA, editor. Biomechanics (structures and systems): a practical approach. New York, NY: IRL Press at Oxford University Press. p 205-231.

  • Fixation
    1. Catch live animal
    2. Narcotize animal to ensure relaxation. 3.75% MgCl2.6H2O in 50% SW; 13oC, 1.5 min (Messenger, J. B., M. Nixon, and K. P. Ryan. 1985. Magnesium chloride as an anesthetic for cephalopods. Comp. Biochem. Physiol. 82C: 203-205.) or 1.5% EtOH in SW for brief period (O'dor, R.K., H.O. Portner, and R.E. Shadwick. 1990. Squid as elite athletes: locomotory, respiratory and circulatory integration. In: Squid as Experimental Animals, D.L. Gilbert, W.J. Adelman, and J.M. Arnold, eds. Plenum Press, New York.) Fix immediately for fully contracted specimen
    3. Dissect out buccal masses
    4. Fix in seawater buffered formalin: 10% v/v.
    5. 100mL Formaldehyde (36%)
    6. 900mL Seawater 35-38 ppm
    7. Let fix for 48 hours (minimum)
    8. Samples may then be transferred to 70% EtOH for storage
  • Dehydrating
    1. Wash briefly in H2O
    2. Transfer to 30% EtOH for 0.5 to 1 hour (Volume of water should be x20 specimen)
    3. Transfer to 50% EtOH for 0.5 to 1 hour
    4. Transfer to 70% EtOH for 0.5 to 1 hour
    5. Transfer to 95% EtOH for 0.5 to 1 hour
    6. Transfer to 100% EtOH for 0.5 to 1 hour
    7. Transfer to 100% EtOH for 0.5 to 1 hour
    8. Transfer to 100% EtOH for 0.5 to 1 hour
    9. Transfer to 50% EtOH/50% Clearing agent for 0.5 to 1 hour
    10. Transfer to 100% Clearing agent for 0.5 to 1 hour
    11. Transfer to 100% Clearing agent for 0.5 to 1 hour
    12. Transfer to 100% Clearing agent for 0.5 to 1 hour
  • Embedding
    1. 1. Transfer specimen to 100% molten paraffin (56 degrees C). (x20 volume, Use vacuum chamber)
    2. 2. Transfer specimen to 100% molten paraffin (56 degrees C).
    3. 3. Transfer specimen to 100% molten paraffin (56 degrees C).
    4. 4. Transfer specimen to 100% molten paraffin (56 degrees C).
    5. 5. Put some paraffin into plastic mold.
    6. 6. Place and orient tissue using hot forceps.
    7. 7. Add more paraffin
    8. 8. Cool rapidly by immersing in water (Cooling slowly creates irregularities and crystals)
  • Cutting
    1. 1. Remove embedded specimen from mold
    2. 2. Trim attachment surface
    3. 3. Use hot spatula to mount embedded specimen to chuck block
    4. 4. Trim block in trapezoidal fashion.
    5. 5. Use rotary microtome with disposable blade holder
  • Staining & mounting
    1. Use staining protocols in Kier, 1992 (Hydrostatic skeletons and muscular hydrostats). Milligan Trichrome/Picro-ponceau with haemotoxylin/Picro Sirius
    2. Mounting medium is non-aqueous. Remove alcohol from slides
      1. Transfer to 50% HemoD/50% EtOH
      2. Transfer to 100% Histoclear
      3. Transfer to 100% Histoclear
    3. 3. Coverslip. Lower slip slowly without trapping bubbles.
    4. 4. Dry.
Cephalopod buccal mass GMA tomography
From Kier WM. 1992. Hydrostatic skeletons and muscular hydrostats, Ch. 9. In: Biewener AA, editor. Biomechanics (structures and systems): a practical approach. New York, NY: IRL Press at Oxford University Press. p 205-231.

  • Fixation
    1. Catch live animal
    2. Narcotize animal. Fix immediately for fully contracted specimen
    3. Dissect out buccal masses
    4. Fix in seawater buffered formalin: 10% v/v.
      1. 100mL Formaldehyde (36%)
      2. 900mL Seawater 35-38 ppm
    5. Let fix for 48 hours (minimum)
    6. Samples may then be transferred to 70% EtOH for storage
  • Semi-thin sectioning using GMA
    Pre-GMA embedding preparation
    1. As the largest blocks that may be embedded measure 1 cm x 1 cm x 4 mm, some beaks must be cut in half or the tissue of interest must be dissected out.
    2. Begin a dehydration series (use 30 mL glass specimen jars and use tissue rotator)
      1. 70% EtOH 1 hr.
      2. 95% EtOH 1 hr.
      (100% EtOH is not needed as GMA is somewhat soluble in water)
    3. Prepare the embedding solutions. (Have used either JB-4 or a Historesin analogue)
      Note: during this step prepare all equipment needed for embedding the tissue.
      Preparing the GMA solution
      1. Using JB-4 mix enough for 4 blocks (4 half beaks)
        1. 100 mL Solution A (use a 100 mL trillium cup)
        2. 0.90 g Catalyst C (weigh on a piece of creased weigh paper)
      2. Add Catalyst C to trillium cup containing Solution A and mix for 20 minutes with a stir bar.
    4. Infiltrate specimen with GMA
      1. Use 30 mL glass specimen jars: 1 jar per piece of tissue
      2. Run through embedding sequence (in each step put into glass vacuum jar and pump down to -1 Atm.
      3. Infiltration sequence:
        1. 10mL JB-4 /10mL 95% EtOH 20 min.
        2. 20mL JB-4 I 1 hr.
        3. 20mL JB-4 II 1 hr.
        4. 20mL JB-4 III 1 hr.
          (Read ahead to Embedding in GMA and assemble equipment during this hour)
        5. 25mL JB-4 Embed 1 hour to complete polymerization
          (Add 1mL of JB-4 Solution B [hardner] to the 25mL in step e)
    5. Embedding in GMA
      1. Get equipment together:
        1. Mold
        2. Backing paper
        3. Water bath and glass
        4. Stopcock grease
        5. Canned air
          (must contain 1,1 difluoroethane [e.g. Dustex 8 or GUST Easy Duster] CAS# 75-37-6. This fluorocarbon does not retard the polymerization process. Do NOT use CAS# 75-45-6 [chlorodifluoromethane] as it is a class II ozone depleting substance and is being phased out. It also does not allow the polymerization to optimally occur.)
        6. Drierite in 5mL vial
        7. Chamber cover
        8. Ice
      2. Label backing paper and stick it onto the mold.
      3. Diagram tissue layout.
      4. Clean the water bath glass and chamber cover.
      5. Apply grease to the chamber cover and connect the canned air.
      6. Fill vial with Drierite and fill water bath with ice water (18 degrees C).
      7. Place glass over water, position chamber cover and insert Drierite.
      8. Organize tissue and load beaks into mold.
        1. Take beak out of JB-4 III
        2. Drag sample through Petrie dish filled with embedding fluid
        3. Orient sample in mold
        4. Use transfer pipette to fill mold with embedding fluid
        5. Do final positioning.
      9. Cover mold/Drierite vial with chamber cover and wait 1 hour to solidify.
      10. Leave the mold in drying oven at 60 degrees C for at least 1 week and preferably longer.
      11. Store tissue blocks at this temperature until sectioning.
    6. Block mounting
      (Some of these directions make references to our specific microtome, the Leica Jung RM2065 motorized microtome, truly the Rolls Royce of microtomes)
      1. Select appropriate aluminum stub.
        1. Remove previous glue residue with a razor blade
        2. Wash with 70% EtOH and dry.
      2. Face the block
        1. Use mill file device and hand file the block side that will be glued to stub.
        2. Cut other side using a long WECK razor. Make the cut parallel to the sanded side.
        3. Glue the block to stub with cyanoacrylate glue (Krazy glue)
        4. Use the facer device with the mill file device to make cut side parallel to stub.
      3. Cut the block profile into a trapezoid shape to cantilever the force of the cutting blade.
        1. Block mounted and edges made parallel. It is ready to be cut.
        2. Trapezoid cut made with long WECK blade.
        3. Direction of blade travel.
          Cut surface trimmed to form another trapezoid cut.
    7. Cutting the block
      1. Turn on machine (Power at back)
      2. Clear distance and slice number
      3. Adjust cutting thickness (3 um)
      4. Move blade all the way back
      5. Designate cutting zone
        1. On downstroke push where cut is to begin.
        2. Move chuck down until past the blade, then push again.
      6. Select 2 or 1 for cutting and then select speed.
      7. Move microscope into view and orient block using adjust knobs and the button.
      8. Begin to cut.
    8. Making slides
      1. Turn on slide warmer to (60 degrees C).
      2. Pull out 10 slides from the acid alcohol cleaning fluid.
      3. Place drops of distilled water onto the first slide. Use a transfer pipette and an Erlenmeyer flask to reduce dust contamination. Use one drop per tissue section and 6 to 8 sections/drops per slide.
      4. Cut sections and float out on water drops using fine forceps or brush.
      5. Place filled slide on warmer and allow to cook for 20 minutes or until dry.
      6. Stain slides after completely dried.
  • Histological staining
    GMA Toluidine blue staining
    1. Make the Toluidine blue stain (2% Toluidine blue in 2% sodium borate)
    2. Load the stain into a 12 cc disposable syringe (Luer lock style) and attach an inline syringe filter.
    3. Preheat slide warmer to 55 degrees - 60 degrees C.
    4. Drip the stain onto the slide once it has warmed up.
    5. Let sit from 25 to 60 seconds (depending on slice thickness)
    6. Rinse thoroughly with distilled water.
    7. Mount coverslips.
  • GMA Trichrome staining
    1. Mordant sections in Bouin's solution for 1 hr. at 56 degrees C or overnight at room temperature (what I do).
    2. Wash in water until yellow colour is removed from the sections.
    3. Rinse in distilled water.
    4. Place slides in Harris' hematoxylin for 30 minutes (check it before using - Changes color in water)
    5. Rinse in water to remove hematoxylin
    6. One quick dip in 1% acid alcohol
    7. Go directly to running tap water to stop reaction.
    8. Dip several times in the lithium carbonate and wash for 5 minutes
    9. Counterstain with Biebrich scarlet-acid fuchsin for 30 minutes
    10. Rinse briefly in distilled water
    11. Counterstain with analine blue for 15 minutes (Masson's acetic analine blue)
    12. Rinse in water, dry and mount.
  • Mounting coverslips
    1. Get some coverslip glass (22 x 60 - 1.5)
    2. Completely dry slides (used canned air)
    3. Immerse in Hemo-D or Histosol or Americlear or, least favorably xylene
      1. Leave for at least 5 minutes
      2. Solvent will go cloudy if contaminated with water
    4. One by one, remove from solvent, put on paper towel and place one or two small drops of Permount onto sections.
    5. Using cover slip, drag drops across sections, then lower cover slip onto slide without trapping any bubbles.
    6. Let dry for 1 + weeks, then remove excess dry Permount with clean razor.
    7. Analyze.

Meiofauna separation
(This technique, which seems to work, was cobbled together through a) a bit of trial and error, b) by reading Sterrer, W. 1971. Smithsonian Contributions to Zoology 76. p. 9-15, and c) several very enjoyable phone calls and emails with Dr. S. Tyler (U. Maine), Dr. E. Rupert (Clemson U.) and Dr. J. Smith, III (Winthrop U.).
  1. Use a PVC corer to get a 10 cm diameter x 40 cm sand core
  2. Divide core into 10 cm long samples and bag
  3. Place one sample into beaker with 250 mL muscle relaxant (7.5% MgCl2.6H2O)
  4. Swirl lightly and let sit 10 minutes
  5. Mix vigorously and filter through a 62 micron mesh-bottomed petri dish
  6. Add another 250 mL of muscle relaxant, swirl and strain
  7. Add 250 mL of fresh seawater, swirl and strain
  8. Fill petri dish lid with fresh seawater and sit mesh-bottomed petri dish into lid. Cover petri dish with another lid and let sit for a while
  9. Examine petri dish and lower cover for specimens
    10. .

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