Team:Bilkent UNAM Turkey/Notebook/Week7

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10/08/2011
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Today we start to making transformation of algae with glass beads method. Protocol is which we follow;
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Transformation of Chlamydomonas with Glass Beads
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contributed by Karen Kindle
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• 1. Grow cells in SGII+NH4NO3 until they reach a density of 1-2 x 10^6/ml.
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• 2. Spin down cells at moderate speed (5000 rpm for 5 min in a GSA type rotor).
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• 3. Resuspend in 1/100 volume SGII+KNO3 and allow to shake at room temperature for 2-4 hours.
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• 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.)
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 +
10/08/2011
 +
<p>Today we start to making transformation of algae with glass beads method. Protocol is which we follow;<p>
-
Note: Several recent experiments suggest that autolysin treatment increases the transformation rate of cw-15 mutants.
+
<div id="container" style="width:100px">
-
• 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.
+
<table border="0">
-
Colonies should be visible in about 6 days.
+
<tr>
-
* 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.
+
<td valign=top>
-
** Use washed agar to remove traces of ammonia and other impurities.
+
<ol><br>
-
***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.
+
<strong><li></strong> Grow cells in SGII+NH4NO3 until they reach a density of 1-2 x 10^6/ml.
 +
<br>
 +
<strong><li></strong> Spin down cells at moderate speed (5000 rpm for 5 min in a GSA type rotor).
 +
<br>
 +
<strong><li></strong> Resuspend in 1/100 volume SGII+KNO3 and allow to shake at room temperature for 2-4 hours.
 +
<br>
 +
<strong><li></strong> 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.)
 +
<br>
 +
<strong>Note:</strong> Several recent experiments suggest that autolysin treatment increases the transformation rate of cw-15 mutants.
 +
<br>
 +
<strong><li></strong>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.
 +
<ul><br>
 +
<li>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.</li>
 +
<br>
 +
<li>Use washed agar to remove traces of ammonia and other impurities.</li>
 +
<br>
 +
<li>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.</li>
 +
<br>
-
Compounds of soluntions;
+
Compounds of soluntions;<br><br>
-
2000 mL SGII-NH4 medium:
+
2000 mL SGII-NH4 medium:<br>
-
NaH2PO4: 7.34 g.
+
NaH2PO4: 7.34 g.<br>
-
K2HPO4: 2.3 g.
+
K2HPO4: 2.3 g.<br>
-
Na acetate (NaC2H3O2 · 3H2O): 4 g.
+
Na acetate (NaC2H3O2 · 3H2O): 4 g.<br>
-
20 mL of 5% Na-citrate.
+
20 mL of 5% Na-citrate.<br>
-
20 mL of 0.1% FeCl3 (must be freshly made).
+
20 mL of 0.1% FeCl3 (must be freshly made).<br>
-
20 mL of 0.4% CaCl2 · 2H2O.
+
20 mL of 0.4% CaCl2 · 2H2O.<br>
-
20 mL of 1.5% MgSO4.
+
20 mL of 1.5% MgSO4.<br>
-
20 mL of 3% NH4NO3.
+
20 mL of 3% NH4NO3.<br>
-
20 mL of trace elements stock.
+
20 mL of trace elements stock.<br>
Divide into appropriate quantities (see Plant Material) and cover
Divide into appropriate quantities (see Plant Material) and cover
with a cotton or foam plug and foil. Autoclave. Reserve 600-700
with a cotton or foam plug and foil. Autoclave. Reserve 600-700
mL for making agar plates. Additional medium will be required if
mL for making agar plates. Additional medium will be required if
-
Part II (optional) is completed.
+
Part II (optional) is completed.<br>
-
1000 mL Trace element stock. One solution containing:
+
1000 mL Trace element stock. One solution containing:<br>
-
H3BO3: 100 mg.
+
H3BO3: 100 mg.<br>
-
ZnSO4 · 7H2O: 100 mg.
+
ZnSO4 · 7H2O: 100 mg.<br>
-
MnSO4 · 4H2O: 40 mg.
+
MnSO4 · 4H2O: 40 mg.<br>
-
CoCl2 · 6H2O: 20 mg.
+
CoCl2 · 6H2O: 20 mg.<br>
-
Na2MoO4 · 2H2O: 20 mg.
+
Na2MoO4 · 2H2O: 20 mg.<br>
-
CuSO4: 4 mg.
+
CuSO4: 4 mg.<br>
25 plates To 700 mL of the SGII-NH4 medium add 10.5 g of agar
25 plates To 700 mL of the SGII-NH4 medium add 10.5 g of agar
(to make 1.5% agar). Autoclave and allow the medium to cool
(to make 1.5% agar). Autoclave and allow the medium to cool
somewhat before pouring. Pour into sterile petri dishes under sterile
somewhat before pouring. Pour into sterile petri dishes under sterile
conditions; if possible let the dishes cool under sterile conditions
conditions; if possible let the dishes cool under sterile conditions
-
before capping to prevent excessive condensation.
+
before capping to prevent excessive condensation.<br>
1000 mL 2X concentration of SGII-NO3 medium. This medium
1000 mL 2X concentration of SGII-NO3 medium. This medium
contains the same components as the recipe for SGII-NH4 medium
contains the same components as the recipe for SGII-NH4 medium
-
but uses KNO3 instead of NH4NO3.
+
but uses KNO3 instead of NH4NO3.<br>
-
NaH2PO4: 7.34 g.
+
NaH2PO4: 7.34 g.<br>
-
K2HPO4: 2.3 g.
+
K2HPO4: 2.3 g.<br>
-
Na acetate (NaC2H3O2 · 3H2O): 4 g.
+
Na acetate (NaC2H3O2 · 3H2O): 4 g.<br>
-
20 mL of 5% Na-citrate.
+
20 mL of 5% Na-citrate.<br>
-
20 mL of 0.1% FeCl3 (must be freshly made).
+
20 mL of 0.1% FeCl3 (must be freshly made).<br>
-
20 mL of 0.4% CaCl2 · 2H2O.
+
20 mL of 0.4% CaCl2 · 2H2O.<br>
-
20 mL of 1.5% MgSO4.
+
20 mL of 1.5% MgSO4.<br>
-
20 mL of 10% KNO3.
+
20 mL of 10% KNO3.<br>
-
20 mL of trace elements stock.
+
20 mL of trace elements stock.<br>
Autoclave. Dilute 250 mL with distilled water to make 500 mL of
Autoclave. Dilute 250 mL with distilled water to make 500 mL of
regular strength SGII-NO4 medium (400 mL for rinsing the agar
regular strength SGII-NO4 medium (400 mL for rinsing the agar
Line 72: Line 488:
and let the medium cool before pouring, then pour into sterile petri
and let the medium cool before pouring, then pour into sterile petri
dishes.Under sterile conditions, rinse the surface of the gelled agar
dishes.Under sterile conditions, rinse the surface of the gelled agar
-
with sterile SGII-NO3 medium.
+
with sterile SGII-NO3 medium.<br>
-
100 mL 20% (wt/vol) polyethylene glycol (MW = 8000).
+
100 mL 20% (wt/vol) polyethylene glycol (MW = 8000).<br>
-
Autoclave.
+
Autoclave.<br>
-
200 mL 70% ethyl alcohol.
+
200 mL 70% ethyl alcohol.<br><br>
-
We are working on obtimization of this protocol.
+
We are working on obtimization of this protocol.<br><br>
-
Fun part is when I look at algae under microscopy it is dancing and really amazing I will try to find a video for you.
+
Funny part is when I look at algae under microscopy it is dancing and really amazing I will try to find a video for you.<br>

Latest revision as of 17:44, 9 July 2011

Team:Bilkent UNAM Turkey/Safety - 2011.igem.org

Team:Bilkent UNAM Turkey/safety

From 2011.igem.org

10/08/2011

Today we start to making transformation of algae with glass beads method. Protocol is which we follow;


  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.

      • Compounds of soluntions;

      2000 mL SGII-NH4 medium:
      NaH2PO4: 7.34 g.
      K2HPO4: 2.3 g.
      Na acetate (NaC2H3O2 · 3H2O): 4 g.
      20 mL of 5% Na-citrate.
      20 mL of 0.1% FeCl3 (must be freshly made).
      20 mL of 0.4% CaCl2 · 2H2O.
      20 mL of 1.5% MgSO4.
      20 mL of 3% NH4NO3.
      20 mL of trace elements stock.
      Divide into appropriate quantities (see Plant Material) and cover with a cotton or foam plug and foil. Autoclave. Reserve 600-700 mL for making agar plates. Additional medium will be required if Part II (optional) is completed.
      1000 mL Trace element stock. One solution containing:
      H3BO3: 100 mg.
      ZnSO4 · 7H2O: 100 mg.
      MnSO4 · 4H2O: 40 mg.
      CoCl2 · 6H2O: 20 mg.
      Na2MoO4 · 2H2O: 20 mg.
      CuSO4: 4 mg.
      25 plates To 700 mL of the SGII-NH4 medium add 10.5 g of agar (to make 1.5% agar). Autoclave and allow the medium to cool somewhat before pouring. Pour into sterile petri dishes under sterile conditions; if possible let the dishes cool under sterile conditions before capping to prevent excessive condensation.
      1000 mL 2X concentration of SGII-NO3 medium. This medium contains the same components as the recipe for SGII-NH4 medium but uses KNO3 instead of NH4NO3.
      NaH2PO4: 7.34 g.
      K2HPO4: 2.3 g.
      Na acetate (NaC2H3O2 · 3H2O): 4 g.
      20 mL of 5% Na-citrate.
      20 mL of 0.1% FeCl3 (must be freshly made).
      20 mL of 0.4% CaCl2 · 2H2O.
      20 mL of 1.5% MgSO4.
      20 mL of 10% KNO3.
      20 mL of trace elements stock.
      Autoclave. Dilute 250 mL with distilled water to make 500 mL of regular strength SGII-NO4 medium (400 mL for rinsing the agar plates and 100 mL for resuspending cells). Use the remaining 750 mL of 2X concentration medium to make agar plates. Additional medium will be required if Part II is done. 50 plates SGII-NO3 medium and 1.5% agar. Rinse the agar beforehand to remove ammonium by placing 7.5 g agar in each of three graduated bottles and bringing to 500 mL with distilled water. After the agar has settled, replace the water and shake; repeat once more, then bring to 250 mL with distilled water. Finally, add sufficient 2X SGII-NO3 medium to bring to 500 mL. Autoclave and let the medium cool before pouring, then pour into sterile petri dishes.Under sterile conditions, rinse the surface of the gelled agar with sterile SGII-NO3 medium.
      100 mL 20% (wt/vol) polyethylene glycol (MW = 8000).
      Autoclave.
      200 mL 70% ethyl alcohol.

      We are working on obtimization of this protocol.

      Funny part is when I look at algae under microscopy it is dancing and really amazing I will try to find a video for you.

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