Endnote Cheet Sheet

EndNote Cheet Sheet

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Arabidopsis Seed Collection

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Arabidopsis Seed Collection

Luca Comai has devised a simple seed collector suitable for high density use (many plants in a small space). It is effective, very cheap, and easy to construct. The instructions for constructing it are given below. They may sound complicated but the collector is really very simple. Feel free to ask questions or send comments.

 

Seed collector II

Introduction

This seed collector is suited for harvesting seed from individual plants grown in small pots (e.g.: square, 6 cm wide at top, tapered to 4.5 cm at bottom, 8 cm high). Each pot can be placed next to other pots to achieve high density spacing.

 

Materials

  1. Adhesive tape, such as VWR or Time-Med pressure sensitive labeling tape 1.8 cm wide. The tape type is not important as long as it sticks and it holds up to greenhouse conditions
  2. Paper stapler
  3. Plastic film, such as Mylar overhead transparency film (0.002 mil, Vu-Color ). The choice of this film is based on characteristics and availability. We recycle the film used for lecturing. Other types maybe suitable: experimenting is the best way to find out whether the film has the right flexibility. Mylar has one disadvantage: it builds an electrostatic charge that attracts seeds. However, in our case, the film comes for free and it looks pretty with all the lecture notes. We prefer instructors who use multicolored pens.

Construction

  1. Cut sheets 12 cm x 42 cm. Roll them lengthwise on a dowel 33 mm in diameter and 50 cm long. The long sides will overlap by about 1.3 cm
  2. Tape the resulting plastic tube once, at one third the distance from one extremity. Make a continuous ring of tape for maximum strength
  3. Flatten and fold back the end of the tube most distant from the tape, in such a way that the seam is central and internal to the fold. Staple the sides just above the fold. The fold line should be 2 cm from the end
  4. About 9 cm above the fold, make a cut perpendicular to the tube. The cut will comprise half or slightly less than half of the circumference and will place the seam in the center of the cut. The collector is finished
  5. Appress the flattened end of the collector to the side of the pot. Place the cut about 5 cm above the rim of the pot and facing the pot. Tape the collector to the pot with a full ring of tape
  6. Gently spread the cut and introduce the young inflorescence. As secondary inflorescences are produced guide them in the collector or remove them
  7. A plant with a fully developed inflorescence can be easily fitted with a collector.
    • Gently lay the pot on its side so that the inflorescence fits over half a sheet of standard printer paper (7.5 cm x 26 cm)
    • Roll the paper to enclose the inflorescence. Make the roll’s diameter smaller than the collector’s. Tape the roll to avoid unfolding and pull it to where the length of the inflorescence-paper roll assembly is longer than that of the collector
    • Place the pot at the edge of a table with the inflorescence leaning out. Gently push the rolled inflorescence into the collector. Once fully inserted, the tip of the paper roll should stick out or be easily reached. Pull the paper roll out, leaving the inflorescence in the collector
    • Tape the collector as in “5”

Comments

The dimension of the collector can be changed to fit any square pot. Its design can also be modified to fit special situations. We harvest the seed by cutting the inflorescence at its base, throwing the label into the collector and closing its ends. Collectors can be washed and reassembled or thrown away. I would appreciate knowing of any improvement. For big pots and when space is not a limitation, our previous seed collector (see Compleat guide, AATDB) works well.

Dna Extraction Procedure From Human Blood

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1. Draw 5 ml of blood using lavender-top Vacutainer (Beckton-Dickinson; EDTA anticoagulant)
2. Keep cool until prep is performed (e.g. in an ice chest), but DO NOT FREEZE. Highest yield will be achieved by extracting within 24 hours.
3. Empty blood into a 15 ml Falcon tube; add 10 ml of Red Cell Lysis Buffer (RCLB) and mix completely by inversion.
RCLB: 1 mM NH4HCO3
115 mM NH4Cl
Note: You can bring pre-weighed NH4Cl and a 1M solution of NH4HCO3, and use the cleanest water available to make RCLB on-site. Try to use distilled water, but in a pinch I suspect that water from a personal filter (e.g. Pur, etc.) would work. If it is to be used immediately, this solution does not need to be autoclaved; if you plan to store it for more than a day, autoclave.
4. Spin 10 minutes @ 1,200g in a clinical centrifuge.
5. Discard supernatant and resuspend cell pellet in 10 ml RCLB; repeat step 4.
6. Discard supernatant; resuspend cell pellet (it should be white now) in 1.8 ml of White Cell Lysis Buffer (WCLB). It should be extremely gelatinous – like snot.
WCLB: 100 mM Tris-Cl (pH 7.6)
40 mM EDTA (pH 8.0)
50 mM NaCl
0.2% SDS
0.05% Sodium azide
Note: This solution (before the addition of SDS) definitely needs to be autoclaved – this means bringing a heavy, well-protected bottle with you, or being completely certain that you will be able to autoclave on site. Add SDS after the autoclaved solution cools. Remember that the DNA is going to sit around in this stuff under less-than-ideal conditions for a considerable period of time…
7. Store each white cell lysate in a 2 ml screw-top tube. DNA should be stable in this form for several weeks at room temperature, although refrigeration is recommended just to be safe.
8. To perform final extraction, add 150 µl of saturated NaCl (~ 6M NaCl) to 400 µl of white cell lysate.
9. Vortex and invert to mix; place on ice for 10′.
12. Centrifuge 5′ @ 12,000 RPM.
12. Add the supernatant to a 1.5 ml tube (550 µl); add 1000 µl absolute (100%) ethanol. Mix by inversion – the DNA precipitate should be visible at this point.
13. Spin 5′ @ 12,000 RPM; discard supernatant.
14. Wash w/ 1 ml 70% ethanol; air dry and resuspend in 100 µl of dilute TE (1:4 w/ H2O). Leave overnight at 4°C to resuspend completely. Check concentration and dilute to 100 ng/µl for stock. Freeze white cell lysates at -70°C for long-term storage.
Recommended field equipment for DNA extraction
1. Clinical centrifuge capable of at least 1,000 RPM, or hand-cranked model
2. 10 ml pipettes (minimum of 1 per extraction session – e.g. 1/day)
3. 15 ml Falcon tubes (1 per sample)
4. P-1000 Pipetteman
5. 1000 µl pipette tips (minimum of 1/sample)
6. 2 ml screw-top tubes
7. Gloves
8. Tube racks (for Eppendorfs and 15 ml tubes)
9. Blood collecting supplies (Vacutainers, needles [21 gauge if possible], plastic housings, tourniquets, alcohol preps, cotton, Band-Aids, biohazard bin for needles)
10. Ice chest (pack the other supplies inside for travel – and bring documentation for customs in case you are asked about all of those needles, white powders and solutions inside)

Inmproving Infection Efficiency Via Virus Centrifugation

1. Ultracentrifuge the virus at 50,000 x g for 90 min at 4°C.
Remove the supernatant. Drain carefully and well (preferably with a pipet) since the viral pellet is glassy and will be a filmy smear on the side of the tube.
 
2. Resuspend the virus to 0.5–1% of the original volume in TNE and incubate overnight at 4oC. Swirling during incubation may damage the virus. Gently pipet the solution to mix only after the overnight incubation, to allow diffusion of the virus. TNE is Tris buffer with NaCl and EDTA, which helps maintain the pH and is appropriate for storage if desired. Media can be used if the virus will be used immediately.
 
3. If desired, perform a second round of ultracentrifugation (Steps 1–2) by pooling previously concentrated virus.
 
4. This step is necessary only if injecting into animals, not if infecting cells in culture: Remove cellular debris and aggregated virus by low speed centrifugation (500 x g) for 5 min at 4oC.
 
5. Determine the viral titers of pre- and post-concentrated viral supernatants.
 
6. Infect target cells according to the Retroviral Gene Transfer and Expression

Immunofluorescence Double Staining Protocol Parallel Approach

1. Preparation of Slides

 

 A. Cell Lines

  • Grow cultured cells on sterile glass cover slips or slides overnight at 37 º C

  •  Wash briefly with PBS
  • Fix as desired. Possible procedures include:

    10 minutes with 10% formalin in PBS (keep wet)

    5 minutes with ice cold methanol, allow to air dry

    5 minutes with ice cold acetone, allow to air dry

  • Wash in PBS

 B. FrozenSections

 

  • Snap frozen fresh tissues in liquid nitrogen or isopentane pre-cooled in liquid nitrogen, embedded in OCT compound in cryomolds. Store frozen blocks at – 80 ºC.

  • Cut 4-8 um thick cryostat sections and mount on superfrost plus slides or gelatin coated slides. Store slides at – 80 ºC until needed.
  •  Before staining, warm slides at room temperature for 30 minutes and fix in ice cold acetone for 5 minutes. Air dry for 30 minutes.
  •  Wash in PBS

 C. Paraffin Sections

  • Deparaffinize sections in xylene, 2x5min.

  • Hydrate with 100% ethanol, 2x3min.
  •  Hydrate with 95% ethanol, 1min.
  •  Rinse in distilled water.
  •  Follow procedure for pretreatment as required.

2. Pretreatments of Tissue Sections

 

Antigenic determinants masked by formalin-fixation and paraffin-embedding often may be exposed by epitope umasking, enzymatic digestion or saponin, etc. Do not use this pretreatment with frozen sections or cultured cells that are not paraffin-embedded.

 

3. Procedure

 

Note: prior to perform double labeling, it is important to test each primary antibody individually and select the best pretreatment(s) for each antibody. It will be ideal if the two primary antibodies require same pretreatment. Otherwise, one should do a further test by treating sections with both pretreatments and then immunostain for each antibody individually. If both antibodies survive the “double pretreatments”, you are ready for immunohistochemistry double staining. Another alternative is to do pretreatments separately for each antibody staining.

  1.  Rinse Sections in PBS-Tween 20 for 2×2 min.

  2. Serum Blocking: incubate sections in normal serum blocking solution – species same as secondary antibody (for example: primary antibodies are mouse and rabbit, and secondary antibodies are horse anti-mouse, and goat anti-rabbit, so horse and goat serum block should be used).
  3. Primary Antibodies: incubate sections in the mixture of two primary antibodies (mouse and rabbit) at appropriate dilution in antibody dilution buffer for 1 hour at room temperature.
  4. Rinse in PBS-Tween 20 for 3×2 min.
  5. Secondary Antibodies: incubate sections in the mixture of two fluorescent conjugated secondary antibodies (FITC conjugated Horse anti-Mouse and Texas Red conjugated Goat anti-Rabbit) in PBS for 30 minutes at room temperature).
  6. Rinse in PBS-Tween 20 for 3×2 min.
  7. Counterstain with DAPI if desired for 20 minutes at room temperature.
  8. Rinse in PBS-Tween 20 for 3×2 min.
  9. Coverslip with anti-fade fluorescent mounting medium and seal with nail polish.
  10. Store slides in dark at 4 °C.

 4. Results:

  •  1st Primary Antibody Staining Sites ——————- green
  •  2nd Primary Antibody Staining Sites——————- red  
  • Double Staining Sites ———————————– yellow
  • Counterstained Nuclei ———————————- light blue

Protocol For Cryosectioning

While the timing of the various steps in this protocol are probably not critical, I tend to prefer to process the tissue all at once to ensure that RNA and/or proteins do not get degraded.

 

Solutions

20% Paraformaldehyde/4% Paraformaldehyde-PBS

200 g paraformaldehyde

1 ml 10N NaOH

up to 1 liter with Q, heat to 65° to dissolve, aliquote and store at -20°.

Mix 100 ml 20% Paraformaldehyde with 50 ml 10X PBS and bring up to 500 ml with Q

filter, and store at 4° for up to 2 weeks

 

Sucrose/PBS

30% sucrose 150 g sucrose

up to 500 ml with 1X PBS

filter sterilize and store at room temperature

 

 

Procedure

• Dissect and fix the tissue in fresh 4% paraformaldehyde on ice for 5-10 minutes.

• Wash for 5 minutes in 1X PBS and repeat.

• Transfer to 30% sucrose until the tissue sinks (5-10 minutes depending on the size of the tissue).

• Transfer through a 1:1 mixture of OCT:sucrose and then into OCT.

• Place the tissue in the cryomold, overlay with OCT, orient and freeze quickly on dry ice.

• Once the tissue is in the mold with OCT it should be oriented and frozen quickly because a film can form on the top of the mold (where the OCT is exposed to air) and make moving the tissue difficult.

• If the size of the mold is small enough place each block into an eppendorf tube and store at -80°C.

• Cut 20 micron sections and place on silinized superfrost slides (Protocol S.6).

• For best results, proceed immediately with immunohistochemical staining.