Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Lyophilized lipids, obtained from Avanti Polar Lipids, are solubilized in a mixture of 96%Chloroform + 4%MetOH to a final concentration of 20mg/ml. Lipids are stored in glass vials with Teflon-lined caps in the -80C freezer.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.
• Add antioxidant BHT (from a 2mg/ml stock in EtOH) to the lipids to make a final concentration of 2μg/ml in the mixed micelle stock solution.
• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.
• To ensure that residual solvent is completely removed, place the flask in high-vacuum for one hour.
• Add DD-water to the flask (amount of water will define your Lipid concentration), vortex rigorously and collect the lipid-detergent mixture / sonicate.
• Add detergent to solubilize and vortex. In general the mixed micelle solution is clear, but some mixtures may have a milky appearance. Sonicate.
• The lipid-detergent mix can be stored for one week at 4C.

example:

• in a flask:
  • 200 μL of DMPC at in [50 mg/ml] (gives 10 mg of lipids)
  • + EtoH (anti-oxidizer)
  • + 2 mL chloroform
• apply reverse-phase evaporation
• let in high-vacuum for about 1 hour
• add 8 mL of water (gives [1.25 mg/ml] )
• vortex rigorously
• sonicate
• Extract 0.8mL (thus 1mg of Lipids)
• sonicate about 20 seconds
• add detergent until solubilization:
  • add 50uL of DDM [10 mg/mL] , then vortex - solution milky
  • add again 50uL of DDM, then vortex - solution is clear. (gives 100 μL of DDM, that is 1mg)
• sonicate
• Add 100 μL of water. Now, we have a 1000uL (with 1mg of lipids and 1mg of DDM).

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3
• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3
• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3

If the protein concentration in the sample exceeds 1mg/ml, dilute the protein to 1mg/ml, preferably by using the same buffer as used for the last steps in the purification protocol. If not possible, use a similar buffer with respect to pH and ionic strength. Care has to be taken not to dilute the detergent in the sample too much so the concentration goes below the CMC, since this may induce aggregation of the protein.

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Tamir's lab protocol to prepare dialysis membrane

1. ​ Cut the tubing by ~10cm a piece, and do about 20 pieces one time
2. ​ Boil the tubing in buffer a for 1hr
3. ​ Wash the tubing with mQ H2O​
4. ​ Boil the tubing in buffer b for 1hr
5. ​ Wash the tubing with mQ H2O
6. ​Boil the tubing in mQ H2O for 1hr
7. ​Wash the tubing with mQ H2O
8. ​ Store the tubing in buffer c at 4ºC

Set up and assembly of the dialysis block

• The dialysis block, including the silicone gaskets should be carefully rinsed with soap and water before the set up of a new dialysis trial. The parts are thereafter sterilized in 70% EtOH, cleansed in DD-water and left to dry. Special care is necessary when rinsing the dialysis wells, since contaminating bacteria may hide there.
• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.

Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Lyophilized lipids, obtained from Avanti Polar Lipids, are solubilized in a mixture of 96%Chloroform + 4%MetOH to a final concentration of 20mg/ml. Lipids are stored in glass vials with Teflon-lined caps in the -80C freezer.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.
• Add antioxidant BHT (from a 2mg/ml stock in EtOH) to the lipids to make a final concentration of 2μg/ml in the mixed micelle stock solution.
• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.
• To ensure that residual solvent is completely removed, place the flask in high-vacuum for one hour.
• Add DD-water to the flask (amount of water will define your Lipid concentration), vortex rigorously and collect the lipid-detergent mixture / sonicate.
• Add detergent to solubilize and vortex. In general the mixed micelle solution is clear, but some mixtures may have a milky appearance. Sonicate.
• The lipid-detergent mix can be stored for one week at 4C.

example:

• in a flask:
  • 200 μL of DMPC at in [50 mg/ml] (gives 10 mg of lipids)
  • + EtoH (anti-oxidizer)
  • + 2 mL chloroform
• apply reverse-phase evaporation
• let in high-vacuum for about 1 hour
• add 8 mL of water (gives [1.25 mg/ml] )
• vortex rigorously
• sonicate
• Extract 0.8mL (thus 1mg of Lipids)
• sonicate about 20 seconds
• add detergent until solubilization:
  • add 50uL of DDM [10 mg/mL] , then vortex - solution milky
  • add again 50uL of DDM, then vortex - solution is clear. (gives 100 μL of DDM, that is 1mg)
• sonicate
• Add 100 μL of water. Now, we have a 1000uL (with 1mg of lipids and 1mg of DDM).

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3
• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3
• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3

If the protein concentration in the sample exceeds 1mg/ml, dilute the protein to 1mg/ml, preferably by using the same buffer as used for the last steps in the purification protocol. If not possible, use a similar buffer with respect to pH and ionic strength. Care has to be taken not to dilute the detergent in the sample too much so the concentration goes below the CMC, since this may induce aggregation of the protein.

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Tamir's lab protocol to prepare dialysis membrane

Steps:

1. ​ Cut the tubing by ~10cm a piece, and do about 20 pieces one time
2. ​ Boil the tubing in buffer a for 1hr
3. ​ Wash the tubing with mQ H2O​
4. ​ Boil the tubing in buffer b for 1hr
5. ​ Wash the tubing with mQ H2O
6. ​Boil the tubing in mQ H2O for 1hr
7. ​Wash the tubing with mQ H2O
8. ​ Store the tubing in buffer c at 4ºC

Set up and assembly of the dialysis block

• The dialysis block, including the silicone gaskets should be carefully rinsed with soap and water before the set up of a new dialysis trial. The parts are thereafter sterilized in 70% EtOH, cleansed in DD-water and left to dry. Special care is necessary when rinsing the dialysis wells, since contaminating bacteria may hide there.
• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.

Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Lyophilized lipids, obtained from Avanti Polar Lipids, are solubilized in a mixture of 96%Chloroform + 4%MetOH to a final concentration of 20mg/ml. Lipids are stored in glass vials with Teflon-lined caps in the -80C freezer.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.
• Add antioxidant BHT (from a 2mg/ml stock in EtOH) to the lipids to make a final concentration of 2μg/ml in the mixed micelle stock solution.
• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.
• To ensure that residual solvent is completely removed, place the flask in high-vacuum for one hour.
• Add DD-water to the flask (amount of water will define your Lipid concentration), vortex rigorously and collect the lipid-detergent mixture / sonicate.
• Add detergent to solubilize and vortex. In general the mixed micelle solution is clear, but some mixtures may have a milky appearance. Sonicate.
• The lipid-detergent mix can be stored for one week at 4C.

example:

• in a flask:
  • 200 μL of DMPC at in [50 mg/ml] (gives 10 mg of lipids)
  • + EtoH (anti-oxidizer)
  • + 2 mL chloroform
• apply reverse-phase evaporation
• let in high-vacuum for about 1 hour
• add 8 mL of water (gives [1.25 mg/ml] )
• vortex rigorously
• sonicate
• Extract 0.8mL (thus 1mg of Lipids)
• sonicate about 20 seconds
• add detergent until solubilization:
  • add 50uL of DDM [10 mg/mL] , then vortex - solution milky
  • add again 50uL of DDM, then vortex - solution is clear. (gives 100 μL of DDM, that is 1mg)
• sonicate
• Add 100 μL of water. Now, we have a 1000uL (with 1mg of lipids and 1mg of DDM).

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3
• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3
• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2
  • 100mM NaCl
  • 0.5mM NaN3

If the protein concentration in the sample exceeds 1mg/ml, dilute the protein to 1mg/ml, preferably by using the same buffer as used for the last steps in the purification protocol. If not possible, use a similar buffer with respect to pH and ionic strength. Care has to be taken not to dilute the detergent in the sample too much so the concentration goes below the CMC, since this may induce aggregation of the protein.

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Set up and assembly of the dialysis block

• The dialysis block, including the silicone gaskets should be carefully rinsed with soap and water before the set up of a new dialysis trial. The parts are thereafter sterilized in 70% EtOH, cleansed in DD-water and left to dry. Special care is necessary when rinsing the dialysis wells, since contaminating bacteria may hide there.
• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.

Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.

• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.
• To ensure that residual solvent is completely removed, place the flask in high-vacuum for one hour.
• Add DD-water to the flask (amount of water will define your Lipid concentration), vortex rigorously and collect the lipid-detergent mixture / sonicate.
• Add detergent to solubilize and vortex. In general the mixed micelle solution is clear, but some mixtures may have a milky appearance. Sonicate.
• The lipid-detergent mix can be stored for one week at 4C.

example:

• in a flask:
  • 200 μL of DMPC at in [50 mg/ml] (gives 10 mg of lipids)

• • + 2 mL chloroform
• apply reverse-phase evaporation
• let in high-vacuum for about 1 hour
• add 8 mL of water (gives [1.25 mg/ml] )
• vortex rigorously
• sonicate
• Extract 0.8mL (thus 1mg of Lipids)
• sonicate about 20 seconds
• add detergent until solubilization:
  • add 50uL of DDM [10 mg/mL] , then vortex - solution milky
  • add again 50uL of DDM, then vortex - solution is clear. (gives 100 μL of DDM, that is 1mg)
• sonicate
• Add 100 μL of water. Now, we have a 1000uL (with 1mg of lipids and 1mg of DDM).

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2

• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2

• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Set up and assembly of the dialysis block

• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.

Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Lyophilized lipids, obtained from Avanti Polar Lipids, are solubilized in a mixture of 96%Chloroform + 4%MetOH to a final concentration of 20mg/ml. Lipids are stored in glass vials with Teflon-lined caps in the -80C freezer.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.
• Add antioxidant BHT (from a 2mg/ml stock in EtOH^?) to the lipids to make a final concentration of 2μg/ml in the mixed micelle stock solution.
• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.

• The lipid-detergent mix can be stored for one week at 4C.

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?
• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?
• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?

If the protein concentration in the sample exceeds 1mg/ml, dilute the protein to 1mg/ml, preferably by using the same buffer as used for the last steps in the purification protocol. If not possible, use a similar buffer with respect to pH and ionic strength. Care has to be taken not to dilute the detergent in the sample too much so the concentration goes below the CMC, since this may induce aggregation of the protein.

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Set up and assembly of the dialysis block

• The dialysis block, including the silicone gaskets should be carefully rinsed with soap and water before the set up of a new dialysis trial. The parts are thereafter sterilized in 70% EtOH^?, cleansed in DD-water and left to dry. Special care is necessary when rinsing the dialysis wells, since contaminating bacteria may hide there.
• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.

Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Lyophilized lipids, obtained from Avanti Polar Lipids, are solubilized in a mixture of 96%Chloroform + 4%MetOH to a final concentration of 20mg/ml. Lipids are stored in glass vials with Teflon-lined caps in the -80C freezer.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.
• Add antioxidant BHT (from a 2mg/ml stock in EtOH^?) to the lipids to make a final concentration of 2μg/ml in the mixed micelle stock solution.
• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.
• To ensure that residual solvent is completely removed, place the flask in high-vacuum for one hour.

• The lipid-detergent mix can be stored for one week at 4C.

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?
• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?
• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?

If the protein concentration in the sample exceeds 1mg/ml, dilute the protein to 1mg/ml, preferably by using the same buffer as used for the last steps in the purification protocol. If not possible, use a similar buffer with respect to pH and ionic strength. Care has to be taken not to dilute the detergent in the sample too much so the concentration goes below the CMC, since this may induce aggregation of the protein.

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Set up and assembly of the dialysis block

• The dialysis block, including the silicone gaskets should be carefully rinsed with soap and water before the set up of a new dialysis trial. The parts are thereafter sterilized in 70% EtOH^?, cleansed in DD-water and left to dry. Special care is necessary when rinsing the dialysis wells, since contaminating bacteria may hide there.
• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.

Protein handling, lipid preparation and setup of dialysis block at NYSBC, by Martin Vink, 12/2008.

Contents

All calculations in the text (under “Set up dialysis block”) are based on the assumption that the protein to be dialyzed arrives at concentration exceeding 1mg/ml. If not, the matrix for the dialysis block setup needs to be re-calculated. This is an important consideration when obtaining proteins from an outside source – ask to obtain the protein at or exceeding a concentration of 1mg/ml.

Protein storage

Pipeline of events for setting up dialysis block

• Prepare lipids up to one week before protein arrival, but preferably as late as possible.
• Wash the dialysis block

Upon protein arrival:

• Analyze the protein sample in a light microscope.
• Run an SDS-PAGE
• Measure protein concentration, unless already determined.
• Measure residual lipid and detergent by TLC.

Just before setting up the dialysis trial:

• Clear the sample by centrifugation.

Most of these events are explained in more detail below.

Protein analysis in light microscope

Procedure: Place a small aliquot of the protein sample (50-100μl) on a microscopy slide or in a clear U-bottomed well in a microtiter plate. Use a dissecting microscope or a microscope for screening 3D crystallization trials. The protein sample is recoverable after analysis.

SDS-PAGE

To assess sample composition and purity, an SDS-PAGE of the protein sample is performed. The SDS-PAGE could also be used for determination of protein concentration (see below).

Choose an appropriate acrylamide concentration for the gel, high for small proteins and low for larger ones. Normally gels with 8-12% acrylamide will do the trick. Don’t forget suitable markers.

Determination of protein concentration

Accurate protein measurement in concentrated membrane protein samples is difficult due to interference with the standard assays (Lowry, Bradford etc.) by lipids and co-concentrated detergent micelles.

There are, however, alternatives:

• An SDS-PAGE can be performed with a known volume of sample and the protein bands can be densitometrically compared to bands of a reference protein ladder (i.e. BSA) by using the program ImageJ^?. A problem is that different proteins bind Coomassie to different extents and therefore have different widths and densities on a gel.
• The absorbance of the protein sample can be measured at 280nm and, by using the extinction coefficient; the protein concentration can be calculated. Check carefully that the buffer does not absorb excessively and use it to blank the spectrophotometer. Estimated extinction coefficients depend on the amounts of aromatics in the protein and can be obtained using the protein parameters tool on the ExPASy^? homepage (http://kr.expasy.org/tools/protparam.html). Paste the sequence of the protein into the box and press “compute parameters” and you will obtain a number of additional parameters including molecular weight, theoretical PI and amino acid composition.

Be sure to always use the same method for determining the protein concentration. All methods have their flaws, but by being consistent more reproducible results are obtained.

TLC

Residual lipid may affect the protein-lipid ratio or it may affect the properties of the lipid bilayer depending on its charge, phase transition temperature, alkyl chain length and saturation. The amount of lipid and detergent in a sample can be estimated by thin-layer chromatography using reference ladders with known lipid and detergent concentrations.

Procedure: see Protocol for TLC of lipids and detergents

Ultracentrifugation

Removal of small precipitates and other contaminating particles before dialysis is done by ultracentrifugation of the sample at 100,000g for 30 minutes.

Choice of detergent

If a detergent screen has not yet been performed, the lipid-detergent stock should be prepared with the detergent in which the protein has been purified.

Lipid preparation

It is important to make the lipid-detergent stocks in a consistent way so that differences in crystallization trial outcomes between different screens cannot be attributed to the mixed micelle preparation.

Lyophilized lipids, obtained from Avanti Polar Lipids, are solubilized in a mixture of 96%Chloroform + 4%MetOH to a final concentration of 20mg/ml. Lipids are stored in glass vials with Teflon-lined caps in the -80C freezer.

Before dialysis, stocks of lipids solubilized in detergent (mixed micelles) are prepared. The ratio lipid:detergent is generally set to 1:2 (2.5mg/ml lipid in 5 mg/ml detergent) but can be changed to meet other criteria.

Procedure:

• Calculate the amount of lipid(s) needed for the stock solution and rapidly, using a pipette, transfer the amount into a small round-bottomed boiling flask.
• Add antioxidant BHT (from a 2mg/ml stock in EtOH^?) to the lipids to make a final concentration of 2μg/ml in the mixed micelle stock solution.
• Add some additional pure chloroform (0.5-1ml) to the flask to ensure that the lipid film produced will be as thin as possible.
• Fit the boiling flask into a rotary evaporator, apply vacuum and turn on the rotator. When all chloroform has been removed, a thin lipid film should be seen in the flask.
• To ensure that residual solvent is completely removed, place the flask in high-vacuum for one hour.
• Add detergent solubilized in DD-water to the flask, vortex rigorously and collect the lipid-detergent mixture. In general the mixed micelle solution is clear, but some mixtures may have a milky appearance.
• The lipid-detergent mix can be stored for one week at 4C.

Preparation of samples for the dialysis block

For the first screen of any given protein, the effect of the following three parameters on 2D crystal formation is evaluated:

• pH (6.0, 7.0 and 8.0)
• LPR (0.1, 0.25, 0.5, 1.0 and 1.5)
• Type of lipid (DMPC, DOPC, POPC, DOPG and E. Coli polar lipids)

In total 3(pH) x 5(LPRs) x 5 (lipids) = 75 crystallization conditions are initially screened for each protein.

Since all lipids are tested in (75/5 =) 15 wells per block, and each lipid is used at 5 different LPRs (see below for specific volumes), each block requires (3x40.5μl ~) 150μl lipid/detergent mix.

Three different dialysis buffers are prepared, varying only in their pH:

• Buffer pH6:
  • 20mM MES pH6.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?
• Buffer pH7
  • 20mM TES pH7.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?
• Buffer pH8
  • 20mM TES pH8.0
  • 5mM MgCl2
  • 100mM NaCl^?
  • 0.5mM NaN3^?

If the protein concentration in the sample exceeds 1mg/ml, dilute the protein to 1mg/ml, preferably by using the same buffer as used for the last steps in the purification protocol. If not possible, use a similar buffer with respect to pH and ionic strength. Care has to be taken not to dilute the detergent in the sample too much so the concentration goes below the CMC, since this may induce aggregation of the protein.

60μl is prepared of each sample for dialysis block. It is most convenient to use a 96-well microtiter plate for sample mixing, since the samples can be prepared according to the matrix to be used in the dialysis block. Extensive labeling of tubes is also avoided. The following amounts of lipids and proteins and the corresponding LPRs (weight to weight) are used for 60μl sample:

Table: Each row of the table is a line containing of one or more cells. Each cell starts and ends with a vertical bar '|'. Any spaces at the beginning of a line are ignored.

Once mixed, the samples should be transferred as quickly as possible to the dialysis block to commence dialysis.

Depending on the outcome of this first screen, parameters are fine tuned for the next one.

Set up and assembly of the dialysis block

• The dialysis block, including the silicone gaskets should be carefully rinsed with soap and water before the set up of a new dialysis trial. The parts are thereafter sterilized in 70% EtOH^?, cleansed in DD-water and left to dry. Special care is necessary when rinsing the dialysis wells, since contaminating bacteria may hide there.
• The silicone gaskets should be attached to the dialysis block before loading the samples into the dialysis wells. It is important to carefully adjust the holes in the gaskets so that they match with the corresponding holes in the block.
• The most critical step in setting up a dialysis screen is securing the two halves of the block together and therefore two people should work together during this part of the protocol: One person is responsible for lowering the upper part of the block, by sliding it along the guide pins until a tight connection is obtained with the lower part. As soon as this is achieved, the other person swiftly secures the screws that hold the two parts together. Speed is instrumental, since even a little wiggling between the two parts of the block by the person trying to keep the two halves together, may cause leakage out of the wells due to capillary action along the dialysis membrane.