Bradford Assay - Revision history

Dicksonb at 14:06, 24 May 2022

2022-05-24T14:06:36Z

← Older revision		Revision as of 14:06, 24 May 2022
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	# Collect lysate or supernatant containing protein. Keep on ice.		# Collect lysate or supernatant containing protein. Keep on ice.
	# Label eight 1.5 mL tubes as: 1, 2, 3, 4, 5, 6, 7, 8. Set up the BSA standard by following '''Table 1'''. (Use the Bio-Rad 2 mg/mL stock. Do not make BSA yourself)		# Label eight 1.5 mL tubes as: 1, 2, 3, 4, 5, 6, 7, 8. Set up the BSA standard by following '''Table 1'''. (Use the Bio-Rad 2 mg/mL stock. Do not make BSA yourself)
	#*Make sure the diluent is the same as buffer that the protein of interest is in. For example, if in PBS, use PBS to dilute the standard~~. If in RIPA buffer, use RIPA, etc~~.		#*Make sure the diluent is the same as buffer that the protein of interest is in. For example, if in PBS, use PBS to dilute the standard.
	#*Mix by pipetting up and down. Avoid creating bubbles in the tube		#*Mix by pipetting up and down. Avoid creating bubbles in the tube
	#Set up 500 μL dilutions of your sample of interest. Make dilutions of 1:5, 1:10, and 1:25.		#Set up 500 μL dilutions of your sample of interest. Make dilutions of 1:5, 1:10, and 1:25.

Dicksonb at 13:49, 27 February 2022

2022-02-27T13:49:08Z

← Older revision	Revision as of 13:49, 27 February 2022
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		-

Dicksonb: Huge edit to fix the previous protocol, which was completely wrong.

2022-02-27T13:47:46Z

Huge edit to fix the previous protocol, which was completely wrong.

Show changes

Dicksonb at 23:26, 17 February 2022

2022-02-17T23:26:24Z

← Older revision		Revision as of 23:26, 17 February 2022
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	''For a 96 well plate''		''For a 96 well plate''

	# Using a 0.2 μm filter and 15 mL syringe, filter ~~approximately 4 mL of~~ Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 5 mL Eppendorf tube or 15 mL ~~falcon~~ tube. Let it warm to room temperature.		# Using a 0.2 μm filter and 15 mL syringe, filter Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 1.5 mL Eppendorf tube or 5 mL tube. Let warm to room temperature.
	# Collect lysate or supernatant ~~from the cells~~. Keep on ice.		# Collect lysate or supernatant containing protein. Keep on ice.
	# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions ~~into a total of 600 μL~~. ~~('''NOTE:''' Do not use the 1:2 diluted sample to prepare the 1:10 tube. Use the original, undiluted sample to prepare the 1:10 dilution)~~		# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions.
	# Prepare a 1 mg/mL stock solution of BSA by dissolving BSA in ddH2O. Prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL.		# Prepare a 10 mg/mL stock solution of BSA by dissolving BSA in ddH2O. Prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL.
	# See '''Figure 1''' for a general plate setup. ~~Using the reverse pipetting technique, pipet 160~~ μL of ~~blank,~~ standard~~, and samples into each appropriate~~ well~~. Avoid creating air bubbles~~.		# See '''Figure 1''' for a general plate setup. 80 μL of sample and standard will be used per well.
	# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.		# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.
	# Pour ~~4 mL of~~ filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye ~~into~~ each well. If a multichannel pipette is not available, '''''quickly'''~~'' pipet 40 μL of dye into each well using a P200 pipette~~.		# Pour filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet dye into each well such that the dye is diluted 1/5 (for example, if 80 μL of sample was loaded per well, then 20 μL of dye will be added in each well) If a multichannel pipette is not available, use the micropipette and pipet into each well '''as quickly as possible'''.
	# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.		# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.
	# Take off the lid of the plate, and place it into a plate reader/spectrometer.		# Take off the lid of the plate, and place it into a plate reader/spectrometer.

Akipp4 at 19:29, 18 January 2022

2022-01-18T19:29:04Z

Dicksonb at 14:23, 15 January 2022

2022-01-15T14:23:51Z

← Older revision		Revision as of 14:23, 15 January 2022
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	''Some things to know'':		''Some things to know'':

	* This protocol uses Bio-Rad products		* This protocol assumes the use of bovine serum albumin (BSA) for the standard curve, which has a linear range of 200 μg/mL to 1000 μg/mL within Bio-Rad's protein assay dye.
	* This protocol assumes the use of bovine serum albumin (BSA) for the standard curve, which has a linear range of 200 μg/mL to 1000 μg/mL within Bio-Rad's protein assay dye~~. Linear range may be improved by using other products, but are more expensive than Bio-Rad's~~.
	* If your protein is within a supernatant containing an acid indicator, such as phenol red, this may interfere with the assay. If your protein is within a solution containing phenol red, then ensure that all samples, standards, and blanks are diluted with that media (e.g. Serum free DMEM).		* If your protein is within a supernatant containing an acid indicator, such as phenol red, this may interfere with the assay. If your protein is within a solution containing phenol red, then ensure that all samples, standards, and blanks are diluted with that media (e.g. Serum free DMEM).
	* This protocol assumes all standards, samples, and blanks are run in triplicates.		* Each tube will have extra volume to ensure you have enough for the last replicate and to avoid air bubbles.
	* Each tube will have extra volume to ensure you have enough for the last replicate.
	* Reverse pipetting technique is highly recommended to avoid air bubbles and to ensure accuracy.		* Reverse pipetting technique is highly recommended to avoid air bubbles and to ensure accuracy.
	* The Bradford Dye is a 5X stock solution.

	''For a 96 well plate''		''For a 96 well plate''

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	# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.		# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.
	# Dump 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.		# Dump 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.
	# Incubate for at least 5 minutes at room temperature.		# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.
	# Take off the lid of the plate, and place it into a plate reader/spectrometer.		# Take off the lid of the plate, and place it into a plate reader/spectrometer.
	# Measure absorbance at 595 nm~~. Export results to Excel~~.		# Measure absorbance at 595 nm.
	[[File:Bradford plate layout.png\|thumb\|'''Figure 1.''' General plate layout \|border\|center]]		[[File:Bradford plate layout.png\|thumb\|'''Figure 1.''' General plate layout \|border\|center]]

Dicksonb at 14:21, 15 January 2022

2022-01-15T14:21:41Z

← Older revision		Revision as of 14:21, 15 January 2022
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	# Take off the lid of the plate, and place it into a plate reader/spectrometer.		# Take off the lid of the plate, and place it into a plate reader/spectrometer.
	# Measure absorbance at 595 nm. Export results to Excel.		# Measure absorbance at 595 nm. Export results to Excel.
	[[File:Bradford plate layout.png~~\|left~~\|thumb\|'''Figure 1.''' General plate layout ]]		[[File:Bradford plate layout.png\|thumb\|'''Figure 1.''' General plate layout \|border\|center]]

Dicksonb at 14:20, 15 January 2022

2022-01-15T14:20:54Z

← Older revision		Revision as of 14:20, 15 January 2022
Line 20:		Line 20:
	# Collect lysate or supernatant that which the protein concentration needs to be calculated for. Keep on ice.		# Collect lysate or supernatant that which the protein concentration needs to be calculated for. Keep on ice.
	# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions into a total of 600 μL. ('''NOTE:''' Do not use the 1:2 diluted sample to prepare the 1:10 tube. Use the original, undiluted sample to prepare the 1:10 dilution)		# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions into a total of 600 μL. ('''NOTE:''' Do not use the 1:2 diluted sample to prepare the 1:10 tube. Use the original, undiluted sample to prepare the 1:10 dilution)
	# ~~Using~~ a 2 mg/mL ~~BSA~~ stock solution ~~(Bio-Rad), prepare~~ a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL. ~~See '''Figure''' '''1.'''~~		# Prepare a 1 mg/mL stock solution of BSA by dissolving BSA in ddH2O. Prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL.
	# See '''Figure 2''' for a general plate setup. Using the reverse pipetting technique, pipet 160 μL of blank, standard, and samples into each appropriate well. Avoid creating air bubbles.		# See '''Figure 1''' for a general plate setup. Using the reverse pipetting technique, pipet 160 μL of blank, standard, and samples into each appropriate well. Avoid creating air bubbles.
	# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.		# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.
	# Dump 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.		# Dump 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.
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	# Measure absorbance at 595 nm. Export results to Excel.		# Measure absorbance at 595 nm. Export results to Excel.
	[[File:Bradford plate layout.png\|left\|thumb\|'''Figure 1.''' General plate layout ]]		[[File:Bradford plate layout.png\|left\|thumb\|'''Figure 1.''' General plate layout ]]











	''Analysis using the standard curve''		''Analysis using the standard curve''

			# Export results from spectrometer to Excel.
			# Subtract the average absorbance of each well by the average absorbance of the blank.
			# Plot the known standard concentrations (X-axis; 0.2, 0.4, 0.6, 0.8, and 1.0 mg/mL) with the average absorbance at each concentration (Y-axis). '''Add a line of best fit and display the equation with R<sup>2</sup> value.''' Ideally, the R<sup>2</sup> value should be ≥ 0.99. R<sup>2</sup> < 0.94 curves should not be used.
			# Calculate the average absorbance for the samples of interest by using the equation of the line graph y = mx+b (Absorbance = Slope*Concentration + b) and solve for x.
			# If the calculated concentration falls within the linear range of the standard curve (0.2 - 1.0 mg/mL), then it is accurate. If necessary, determine the concentration of the undiluted sample by multiplying by the dilution factor.

	#		#

	#		#

Dicksonb at 18:47, 14 January 2022

2022-01-14T18:47:17Z

← Older revision		Revision as of 18:47, 14 January 2022
Line 27:		Line 27:
	# Take off the lid of the plate, and place it into a plate reader/spectrometer.		# Take off the lid of the plate, and place it into a plate reader/spectrometer.
	# Measure absorbance at 595 nm. Export results to Excel.		# Measure absorbance at 595 nm. Export results to Excel.
			[[File:Bradford plate layout.png\|left\|thumb\|'''Figure 1.''' General plate layout ]]

	''Analysis using the standard curve''		''Analysis using the standard curve''

Dicksonb at 03:00, 14 January 2022

2022-01-14T03:00:34Z

← Older revision		Revision as of 03:00, 14 January 2022
Line 4:		Line 4:

	'''Protocol'''		'''Protocol'''

			''Some things to know'':

			* This protocol uses Bio-Rad products
			* This protocol assumes the use of bovine serum albumin (BSA) for the standard curve, which has a linear range of 200 μg/mL to 1000 μg/mL within Bio-Rad's protein assay dye. Linear range may be improved by using other products, but are more expensive than Bio-Rad's.
			* If your protein is within a supernatant containing an acid indicator, such as phenol red, this may interfere with the assay. If your protein is within a solution containing phenol red, then ensure that all samples, standards, and blanks are diluted with that media (e.g. Serum free DMEM).
			* This protocol assumes all standards, samples, and blanks are run in triplicates.
			* Each tube will have extra volume to ensure you have enough for the last replicate.
			* Reverse pipetting technique is highly recommended to avoid air bubbles and to ensure accuracy.
			* The Bradford Dye is a 5X stock solution.

			''For a 96 well plate''

			# Using a 0.2 μm filter and 15 mL syringe, filter approximately 4 mL of Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 5 mL Eppendorf tube or 15 mL falcon tube. Let it warm to room temperature.
			# Collect lysate or supernatant that which the protein concentration needs to be calculated for. Keep on ice.
			# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions into a total of 600 μL. ('''NOTE:''' Do not use the 1:2 diluted sample to prepare the 1:10 tube. Use the original, undiluted sample to prepare the 1:10 dilution)
			# Using a 2 mg/mL BSA stock solution (Bio-Rad), prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL. See '''Figure''' '''1.'''
			# See '''Figure 2''' for a general plate setup. Using the reverse pipetting technique, pipet 160 μL of blank, standard, and samples into each appropriate well. Avoid creating air bubbles.
			# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.
			# Dump 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.
			# Incubate for at least 5 minutes at room temperature.
			# Take off the lid of the plate, and place it into a plate reader/spectrometer.
			# Measure absorbance at 595 nm. Export results to Excel.


			''Analysis using the standard curve''

			#

	#		#

← Older revision		Revision as of 19:29, 18 January 2022
Line 2:		Line 2:

	The Bradford assay should be used to quantify total protein within a sample relative to a standard curve. This is important when running proteins on SDS-PAGE/Western blot, where each lane must contain an identical amount of total protein. This way, the intensity of the bands can be compared between different lanes, as standardizing the amount of protein loaded per lane will account for any variation in cell count from which the proteins were derived.		The Bradford assay should be used to quantify total protein within a sample relative to a standard curve. This is important when running proteins on SDS-PAGE/Western blot, where each lane must contain an identical amount of total protein. This way, the intensity of the bands can be compared between different lanes, as standardizing the amount of protein loaded per lane will account for any variation in cell count from which the proteins were derived.
			'''Protocol'''

	'''Protocol'''		'''Protocol'''
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	# Using a 0.2 μm filter and 15 mL syringe, filter approximately 4 mL of Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 5 mL Eppendorf tube or 15 mL falcon tube. Let it warm to room temperature.		# Using a 0.2 μm filter and 15 mL syringe, filter approximately 4 mL of Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 5 mL Eppendorf tube or 15 mL falcon tube. Let it warm to room temperature.
	# Collect lysate or supernatant ~~that which~~ the ~~protein concentration needs to be calculated for~~. Keep on ice.		# Collect lysate or supernatant from the cells. Keep on ice.
	# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions into a total of 600 μL. ('''NOTE:''' Do not use the 1:2 diluted sample to prepare the 1:10 tube. Use the original, undiluted sample to prepare the 1:10 dilution)		# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions into a total of 600 μL. ('''NOTE:''' Do not use the 1:2 diluted sample to prepare the 1:10 tube. Use the original, undiluted sample to prepare the 1:10 dilution)
	# Prepare a 1 mg/mL stock solution of BSA by dissolving BSA in ddH2O. Prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL.		# Prepare a 1 mg/mL stock solution of BSA by dissolving BSA in ddH2O. Prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL.
	# See '''Figure 1''' for a general plate setup. Using the reverse pipetting technique, pipet 160 μL of blank, standard, and samples into each appropriate well. Avoid creating air bubbles.		# See '''Figure 1''' for a general plate setup. Using the reverse pipetting technique, pipet 160 μL of blank, standard, and samples into each appropriate well. Avoid creating air bubbles.
	# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.		# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.
	# ~~Dump~~ 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.		# Pour 4 mL of filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 40 μL of dye into each well. If a multichannel pipette is not available, '''''quickly''''' pipet 40 μL of dye into each well using a P200 pipette.
	# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.		# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.
	# Take off the lid of the plate, and place it into a plate reader/spectrometer.		# Take off the lid of the plate, and place it into a plate reader/spectrometer.