Crystallography‎ > ‎

Protocols


Glycerol stocks

Add 1 ml (or an equal volume) of glycerol solution to 2mls of mid-log culture or 1ml of freshly saturated culture;
mix gently, then freeze rapidly in liquid nitrogen or on dry ice. Store at -20°C and - 80°C.

Glycerol solution (per 100 ml):

65% glycerol (v/v)     65 ml 100% glycerol
0.1M Mg SO4            10 ml 1M Mg SO4
0.025M Tris pH 8.0     2.5 ml 1M Tris
22.5 ml ddH2O

autoclave to sterilize.

Columns regeneration

Ni-NTA column

Quick
1. Wash with ddH2O for 1hr at 3mL/min.
2. Wash with .1M EDTA for 1hr at 3mL/min.
3. Wash with ddH2O for 1hr at 3mL/min.
4. Wash with .1M NiSO4 for 1hr at 3mL/min.
5. Wash with ddH2O for 1hr at 3mL/min.

More detailed
1) dd water for 10min
2) 2 volumes 8M urea
3) 2 volumes water
4) 3 volumes 2% SDS
5) 2 volumes water
6) 5 volumes 0.1MEDTA
7) 2 volumes water
8) 2 volumes 0.1MNiSO4
9) 6 volumes water
10) 2 volumes buffer needed (before using)

Source Q column

1. Wash with ddH2O for 1hr at 3mL/min.
2. Wash with .1M NaOH for 1hr at 3mL/min.
3. Wash with ddH2O for 1hr at 3mL/min.
4. Wash with 1.5M NaCl for 1hr at 3mL/min.
5. Wash with ddH2O for 1hr at 3mL/min.
6. Wash with ddH2O for 1hr at 3mL/min.

TE Buffer

Purpose: solution containing Tris buffer and EDTA in various ratios used commonly as buffer for DNA.
Standard TE Buffer — T10E1 (100ml):

1 ml 1 M Tris (desired pH, typically 8.0) – resulting concentration 10 mM
0.2 ml 0.5 M EDTA
ddH20 to 100ml


autoclave to destroy DNase that may be present
TE may be kept at room temperature or 4°C.

Agarose gel


Weigh 0.24g of agarose in 30 ml of 1x TAE buffer
Warm in microwave oven with 50% power for 1 min
Assemble wells
After agarose cool down, add 1.5 ml of EtBr in 30 ml of gel
Pour into the cast and wait until gel solidfy

TAE (50X)

Used as a buffer when running DNA in agarose gels.

242 g Tris base
57.1 ml glacial acetic acid
37.2 g Na
2EDTA-2H2O
ddH2O to 1 L

Resulting solution when diluted:

40mM Tris-acetate
2 mM EDTA

pH of 1X solution is » 8.5
May be kept at room temperature.

Protein Concentration

Absorbance
Blank 200uL ddH2O
Tube 1 (1:100) 1uL protein + 199uL ddH2O
Tube 2 (1:50) 2uL protein + 198uL ddH2O
Turn UV light on and let it warm up for 10 min, measure absorbance at 280nm.
Conc(mg/mL) = A280*dilution factor


Bradford Method

Add Protein/BSA, then dye, then ddH2O.


Columns 1 and 2 are for the standard curve and 3 and 4 are for the protein conc.
Do Absorbance concentration and make appropriate dilutions first to make sure protein concentration are between 1-10ug/uL. Test two different concentrations of each protein (i.e. 3ug/uL and 6ug/uL). Average two different concentrations(multiplied by dilution factor) to get the final concentration.

Absorbance Assay (280 nm)
Proteins in solution absorb ultraviolet light with absorbance maxima at 280 and 200 nm. Amino acids with aromatic rings are the primary reason for the absorbance peak at 280 nm. Peptide bonds are primarily responsible for the peak at 200 nm. Secondary, tertiary, and quaternary structure all affect absorbance, therefore factors such as pH, ionic strength, etc. can alter the absorbance spectrum.
1. Warm up the UV lamp (about 10-15 min.)
2. Adjust wavelength to 280 nm
3. Calibrate to zero absorbance with buffer solution only
4. Measure absorbance of the protein solution
5. Adjust wavelength to 260 nm
6. Calibrate to zero absorbance with buffer solution only
7. Measure absorbance of the protein solution

Analysis
Unknown proteins or protein mixtures. Use the following formula to roughly estimate protein concentration. Path length for most spectrometers is 1 cm.
Concentration (mg/ml) = Absorbance at 280 nm divided by path length (cm.)
Pure protein of known absorbance coefficient. Use the following formula for a path length of 1 cm. Concentration is in mg/ml, %, or molarity depending on which type coefficient is used.
Concentration = Absorbance at 280 nm divided by absorbance coefficient
To convert units, use these relationships:

Mg protein/ml = % protein divided by 10 = molarity/protein MW

Unknowns with possible nucleic acid contamination. Use the following formula to estimate protein concentration:

Concentration (mg/ml) = (1.55 x A280) - 0.76 x A260)

Absorbance coefficients of some common protein standards:

Bovine serum albumin (BSA): 63
Bovine, human, or rabbit IgG: 138
Chicken ovalbumin: 70

SDS-PAGE

- Separation-gel: (preparation for 2 gels)
%7.5911131518
30% Acrylamide/bis-acrylamide2.42.93.54.14.65.8
1M Tris pH 8.64.84.84.84.84.63.6
H2O2.21.71.10.500
10% SDS0.0950.0950.0950.0950.0950.095
TEMED0.00490.00490.00490.00490.00490.0049
   degass ~5min, add 68ul APS (2 gels)

- Stacking-gel:


2 gels4 gels
30% Acrylamide/bis-acrylamide0.651.3
0.5M Tris pH 6.80.71.4
H2O2.34.6
10% SDS0.056113
TEMED0.0040.008

degass ~5min, add 37.5ul APS (2gels)

- Procedures:

Gel preparation
1. Setup equipment and check leakage using ethanol
2. Prepare separation gel as outlined above
3. Vacuum gel solution for ~10min to get rid of gas
4. Add 136ul (4 gels) ammonium persulfate (stored in –20C)
5. Load separation gel on equipment and add 1ml 1% SDS on the gel top (to prevent oxidation)
6. Prepare stacking get and degass for 5-10 min
7. Add 75ul (4 gels) ammonium persulfate right before loading
8. Dump SDS and load 1ml stacking gel on top
9. When gel is ready, wrap with film and store at 4C (leave comb inside)

Gel running
1. Mix loading dye (2.5ul) and protein solution (7.5ul)
2. Heat the mix at 100 C for 2min
3. Set up equipment; add running buffer (be sure to fill the inside of glasses)
4. Load sample and run
5. When the band reaches bottom, stop

Gel staining
1. Stain gel for 15 min
2. Wash gel with H2O
3. Add dis-stain buffer and shake for 15 min and change buffer
4. Shake overnight

Gel drying
1. Cover gel with film
2. Put in vacuum dryer at 80C for 1hr

Buffer preparation
4X Sample Buffer
4x sample buffer is added into each sample before loading the sample on the SDS- PAGE. e.g. 10 ul of loading volume contain 2.5 ul of 4x sample buffer and 7.5ul of protein sample.
For making 10 ml:
0.5 M Tris pH6.8 5 ml
glycerol (100%) 4 ml
SDS 0.8 g
Bromphenol blue 2 mg
Dissolve all ingredients above, add ddH2O to bring volume to 10ml. Aliquot into eppendorff tubes with 0.5ml per tube. Store in -20°C. Before use, add 50 ul of 1M DTT (1:10 dilution) and 20ul of 14.3M b-mercatptoethanol to each tube. Label the tube as “DTT added”.


- Comassie blue stain:

Isopropanol 300ml
Acetic acid 100ml
H2O 600ml
Coomassie Brilliant Blue 2.5g

SDS-Gel on bacterial samples

1. Take 1 mL of cells from culture and measure the cell density (OD600)
2. Spin down that mL of cells
3. Divide the OD600 by 10 and resuspend cells in that many mL of 1.5 X sample buffer (SB). (i.e. if your OD600 is 1.2, resuspend cells in 120 microliters 1.5 X S.B.)
4. Boil sample immediately for ~5 minutes
5. Store in refrigerator until ready to load. Samples are stable for at least several days (and probably much longer).
6. At first load ~10 microliters of sample per lane. Adjust volume to suit your results.

How to calculate primer concentration or how much in which to resuspend?

Example 1:

The COA specifies we have 24 nmole of oligo. If we resuspended oligo in 1 ml:
1 ml = .001 L
24 nmole/0.001 L = 24000 nmole/L or 24000 nM
24000 nmole/L X 1 umole/1000 nmole = 24 umole/L or 24 uM
Explanation: We convert the volume in which the oligo was resuspended into liters.
Then the total nmole amount of oligo is divided by the volume to get nM conc. The nmoles are converted to umole to get the uM conc.

Example 2:

Making 100 uM primer stock:
If COA specifies we have 24 nmole of oligo:
24 nmole X 1umole/1000nmole = 0.024 umole
0.024 umole/100 umole/liter = 0.00024 L
0.00024 L X 1000 ml/L = 0.24 ml or 240 ul
Explanation: We convert from nmole to umole then divide by the desired concentration in umole/L. 
(The umoles cancel out giving the needed volume in liters)
We then convert liters to ml. So in this example the oligo should be suspended in 0.24
ml to get a 100 uM solution.

Example 3:

Calculate from OD.
If primer OD is 0.14:
If the ug/OD reported on the COA is 36.6:
0.14 OD/ml X 1000ul/10ul = 14 OD/ml stock
14 OD/ml X 36.6 ug/OD = 512.4 ug/ml
Explanation: The OD/ml is multiplied by the dilution factor to get the stock OD/ml.
The OD is converted to ug/ml by multiplying the OD/ml of the stock by the ug/OD
conversion factor listed on the COA. The ug cancel out giving ug/ml.
If the nmole/OD reported on the COA 4.9:
0.14 OD/ml X 1000ul/10ul = 14 OD/ml stock
14 OD/ml X 4.9 nmole/OD = 68.6 nmole/ml
68.6 nmole/ml X 1000 ml/L = 68,600 nmole/L or 68,600 nM
68,600 nmole/L X 1 umole/1000 nmole = 68.6 umole/L or 68.6 uM
Explanation: The OD/ml is multiplied by the dilution factor to get the stock OD/ml.
The OD is converted to nmole using the conversion factor on the COA. Then ml are
converted to liters and nmole are converted to umole to get the uM concentration.


Example 4:

Calculate from MW.
The COA specifies a molecular weight for the oligo of 7440.0:
7440.9 g/mole = 7440.9 ug/umole
7440.9 ug/umole X 68.6 umole/L = 510445.74 ug/L
510445.74 ug/L X 1 L/ 1000 ml = 510.4 ug/ml
Explanation: g/mole is the same as ug/umole. The molecular weight expressed in
ug/umole is multiplied by the uM concentration determined in example 3. The umole
cancel out leaving ug/L. Liters are converted to ml to give the ug/ml concentration.

Mini-prep of plasmid DNA


1. Grow culture in 3 ml of LB/amp for 8-12 hours at 37oC. (do it afternoon and let it go O/N,
harvest the cell next morning)
2. Transfer 1.5 ml of culture to a microcentrifuge tube and spin for 30 seconds.
3. Pour off the supernatant, remove all residual LB.
4. Resuspend the cell pellet by votexing or pipetting in 300 l STET.
5. Add 20-25 l lysozyme (10mg/ml stock in water and kept frozen at -20C).
6. Place tubes in boiling water for 2 min.
7. Spin 5 min. in microcentrifuge.
8. Remove mucoid pellet with a toothpick.
9. Add an equal volume ( usually 250 l ) of 75% isopropanol/25% 10 M ammonium acetate to
the supernatant, mix vigorously, and spin 5 min in the microcentrifuge.
10. Rinse the pellet well with 70% ethanol and dry in a speed Vac. for 5 min.
(or air dry on the bench)
11. Resuspend in 25-50 ml T20RNase buffer that contain RNase and incubate at 37 degrees (C) for 20 min, store at 4 degrees (C)

Midi-prep of plasmid DNA

Preparation:

Autoclave: 2 centrifuge bottles, 2 glass tubes, 3 250ml flasks
1. DNA is transformed into XL 10-gold competent cell and plate it with Amp-LB plate
2. Pick colony and grow 3ml small culture; add Amp to 50ug/ml
3. Grow big culture (150-200ml LB-Amp) in flask

Midiprep

1. Transfer amplified clutures from flasks into large plastic centrifuge bottles
2. Spin 10,000g for 10min at 4 oC
3. Pour off supernatant, blot excess. Resuspend the pellet in 3ml of Cell Resuspension Solution (I)
   Be sure to suspend completely
4. Transfer to large glass centrifuge tubes. Add 3 ml of Cell Lysis Solution (II). Invert several times. Suspension should be clear
   No vortex!
5. Add 3ml Neutralization Solution (III) and mix by inverting tubes (* No vortex)
6. Centrifuge at 14,000g for 20min at 4 oC
7. Label columns and set up vacuum pump
8. Transfer the supernatant into Falcon 50ml tubes
   Be careful don’t get any white precipitate9. Add 10ml resin to DNA Swirl to mix
10. Transfer resin/DNA to Midi-column
11. Wash with 15ml column Wash Solution 2 times. Dry the resin for 30s
    Don’t dry more than 30s12. Set up Eppendorf tubes for spinning; 2 sets, one labled one not; heat TE buffer at 65 oC
13. Cut off reservoir and transfer column to Eppendorf tube. Spin 10,000g for 20s to remove excess alcohol.
14. Transfer column to labeled tube. Elute DNA with 300ul of TE and let it sit for 1min.
15. Spin 10,000g for 30s
16. Determine concentration by UV spec
17. Store at 4 oC