Team:Bilkent UNAM Turkey/Protocols

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<h1> Growth Conditions </h1>
<h1> Growth Conditions </h1>
'''Tris Acetate Phosphate (TAP) medium
'''Tris Acetate Phosphate (TAP) medium
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Revision as of 22:44, 21 September 2011

Growth Conditions

'''Tris Acetate Phosphate (TAP) medium ''' from Gorman, D.S., and R.P. Levine (1965) Proc. Natl. Acad. Sci. USA 54, 1665-1669. This is probably the most widely-used medium at present for experimental work. Following solutions are prepared. 1. TAP salts NH4Cl 15.0 g MgSO4 . 7H2O 4.0 g CaCl2 . 2H2O 2.0 g water to 1 liter 2. phosphate solution K2HPO4 28.8 g KH2PO4 14.4 g water to 100 ml 3. Hutner's trace elements To make the final medium, mix the following: 2.42 g Tris 25 ml solution #1 (salts) 0.375 ml solution #2 (phosphate) 1.0 ml solution #3 (trace elements) 1.0 ml glacial acetic acid water to 1 liter For solid medium, add 15 g agar per liter Autoclave. For Tris-minimal medium omit the acetic acid and titrate the final solution to pH 7.0 with HCl Reference: [[http://www.chlamy.org/TAP.html]] '''Hutner's trace elements''' Hutner et al. (1950) Proc. Am. Philos. Soc. 94, 152-170 This mixture is used both in TAP and in the Sueoka high salt medium. For a detailed analysis of how well this trace elements solution meets the nutritional requirements of C. reinhardtii, see Merchant et al. (2006) Biochim. Biophys. Acta 1763, 578-594. For 1 liter final mix, dissolve each compound in the volume of water indicated. The EDTA should be dissolved in boiling water, and the FeSO4 should be prepared last to avoid oxidation. compound amount water EDTA disodium salt 50 g 250 ml ZnSO4 . 7 H2O 22 g 100 ml H3BO3 11.4 g 200 ml MnCl2 . 4 H2O 5.06 g 50 ml CoCl2. 6 H2O 1.61 g 50 ml CuSO4 . 5 H2O 1.57 g 50 ml (NH4)6Mo7O24. 4 H2O 1.10 g 50 ml FeSO4. 7 H2O 4.99 g 50 ml Mix all solutions except EDTA. Bring to boil, then add EDTA solution. The mixture should turn green. When everything is dissolved, cool to 70 degrees C. Keeping temperature at 70, add 85 ml hot 20% KOH solution (20 grams / 100 ml final volume). Do NOT use NaOH to adjust the pH. Bring the final solution to 1 liter total volume. It should be clear green initially. Stopper the flask with a cotton plug and let it stand for 1-2 weeks, shaking it once a day. The solution should eventually turn purple and leave a rust-brown precipitate, which can be removed by filtering through two layers of Whatman#1 filter paper, repeating the filtration if necessary until the solution is clear. Store refrigerated or frozen convenient aliquots. Some people shorten the time for formation of the precipiate by bubbling the solution with filtered air. If no precipitate forms, the solution is still usable. However, you might want to check the pH in this case and adjust it to around 7.0 using either KOH or HCl as needed. To prepare sulfur-free trace elements for hydrogen generation, the sulfate salts can be replaced with equimolar chloride salts (ZnCl2 10.0 g; CuCl2 . 2 H2O 1.00 g; FeCl2 . 4 H2O, 3.60 g). . Reference: [[http://www.chlamy.org/trace.html]]

Transformation of Chlamydomonas with Glass Beads

contributed by Karen Kindle

  • 1. Grow cells in SGII+NH4NO3 until they reach a density of 1-2 x 10^6/ml.
  • 2. Spin down cells at moderate speed (5000 rpm for 5 min in a GSA type rotor).
  • 3. Resuspend in 1/100 volume SGII+KNO3 and allow to shake at room temperature for 2-4 hours.
  • 4. If desired, treat cells with gamete autolysin (prepared as described in The Chlamydomonas Sourcebook*) for approximately 45 minutes. I usually resuspend the cells in about 1/25 the original volume in autolysin. Spin and wash once with SGII+KNO3 before transformation. Optional: Test effectiveness of lysis by sensitivity to 0.05% NP-40. (Count duplicate samples +/- detergent in hemacytometer.)

Note: Several recent experiments suggest that autolysin treatment increases the transformation rate of cw-15 mutants.

  • 5. Work in lots of a few tubes at a time. Add 300 microliters cells, 100 microliters 20% polyethylene glycol, 1-2 micrograms DNA (linearized DNA generally transforms somewhat more efficiently than supercoiled), and 300 mg sterile glass*** beads. Vortex for 15-30 seconds at top speed on a Fisher Vortex Genie 2 mixer; plate cells immediately on an SGII+KNO3 selective agar**. Depending on age and moisture of plates, parafilm immediately or after 24 hours; and put in the light.

Colonies should be visible in about 6 days.

* The protocol is followed except that the culture supernatant of the mated gametes is filtered through a 0.22-0.45 micrometer nitrocellulose filter. I don't know how much of the activity (if any) is lost in this protocol, but the requirement for sterility to my mind outweighs any loss in this step. After the filtration, the supernatant is frozen in 50 ml lots at -20 deg C.

** Use washed agar to remove traces of ammonia and other impurities.

***Glass beads are approximately 0.5 mm in diameter and are available from Thomas. We wash them with acid, rinse them with water until the pH is neutral, dry them, and then weigh 300 mg into glass culture tubes and bake for approximately 2 hours at 400 deg F.

 

Different Protocol to Transform Chlamy with Glass Beads.

1. Grow cells in appropriate medium (permissive) to a density of ~2 x 106/ mL.

2. Collect cells by centrifugation in sterile centrifugation bottles at room temperature (3500 g x 10 min). Discard supernatant.

3. Resuspend cells in 1/25 – 1/50 initial volume in selective medium with a cotton-plugged pipet. Transfer to a sterile centrifugation tube.

4. Collect cells by centrifugation at room temperature (3500 g x 10 min). Discard supernatant

5. Resuspend cells at approximately in 1/70 initial volume in selective medium (approximately 3.0 x 108 cells / mL.

Count a 1/100 dilution with the hemacytometer under the microscope. Adjust the volume to obtain a concentration of 2. x 108 cells / mL.

6. To a tube containing 0.3g glass beads (sterilized by baking) add:

- 0.3 mL cell suspension.

- ~ 0.5 – 1.0 ug DNA.

7. Vortex at full speed for 15 seconds.

 

8. Pour the contents of the tube on a selective plate, gently tilt and rotate the plate to spread the medium evenly.

9. Allow the liquid to dry (protect from light to avoid phototactic movements of the cells). Seal the plates with Parafilm (Micropore tape for minimal medium), and incubate under appropriate conditions for selection. (If cells were grown under auxotrophic conditions (acetate, dark) and will be selected for photoautotrophic growth (minimal, light), put the plates in dim light for 16 – 24 hours before transferring to light. Colonies will appear within 1-3 weeks depending on the selection applied.)

References:

Kindle, K. (1990) High-frequency nuclear transformation of Chlamydomonas reinhardtii. Proc. Natl. Acad. USA 87: 1228-1232.

[[http://www.ncbi.nlm.nih.gov/pmc/articles/PMC53444/]]

B. Electroporation method for nuclear transformation of Chlamydomonas reinhardtii

Materials

- Cell-wall deficient host cell strain

- Sterile centrifugation bottles and tubes

- Electroporation cuvettes

- Plates with appropriate solid medium for selection of the transformants

- DNA with selection marker

- TAP, 40mM sucrose

- TAP, 40mM sucrose, 0.4% PEG 8 000 - Starch 20%

Starch 20% preparation

20 g starch in a centrifuge tube

Wash with ethanol 100%

Wash with water

Repeat 2 times

Resuspend in 100 ml Ethanol 70%

Aliquots of 20 ml and keep at room temperature

The day of transformation, centrifuge an aliquot 1 minute at 1 000 rpm

Wash 4 times with TAP + sucrose 40 mM

Resuspend in 20 ml of TAP + sucrose 40 mM + PEG 8 000 0.4%

References.

Boynton et al (1988) Chloroplast transformation in Chlamydomonas with high velocity microprojectiles. Science 240: 1534-1538.

Finer et al. (1992) Development of the particle inflow gun for DNA delivery to plant cells. Plant Cell Reports 11: 323-328.















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