Hello everyone, I am working with a 25 kDa protein which is 6xHis tagged. I have the gene coding for the protein in pET15b and expression is being done in BL 21 DE3. I tried purification and had more than one band appearing on my gel. In addition to a band of the expected size, I am having more bands larger molecular weights. I am using the Ni-NTA Superflow matrix from Qiagen and the buffers used are as follows:
Lysis
20mM HEPES, 200mM NaCl, 1mM MgCl2, 5mM 2 Mercapto-ethanol with protease inhibitors.
Wash 1
20mM HEPES, 10mM imidazole and 0.5M NaCl
Wash 2
20mM HEPES, 20mM imidazole and 0.5M NaCl
Elution
20mM HEPES, 200mM NaCl and 500mM imidazole.
I analysed both the pellet and supernatant after the lysis but there wasn't any band of interest appearing in the pellet.
Thank you in advance for your responses.
As Stefan mentioned, adding a small amount of imidazole to your lysis/binding buffer is recommended by most Ni-NTA column/beads manufacturers. The exact amount varies from protein to protein, but general 10-20 mM imidazole (even 30 mM) is tolerated by most proteins and it greatly helps to reduce non-specific binding.
Also, high NaCl concentrations help with this problem. You use 500 mM in your wash and elution buffers, but only 200 mM in your lysis/binding buffer. I always use 500 mM in all the buffers and, for some proteins that are highly charged (i.e. proteins with pI > 9) I even use 1 M NaCl.
Some of the contaminating proteins could actually be bound to your protein rather than to the Ni-NTA matrix. Adding a small amount of a reducing agent will help with this. For Ni-NTA I use a maximum of 3 mM betamercaptoethanol (BME) in my buffers, higher concentrations of BME or using DTT instead of BME will react with your Ni-NTA matrix and produce a brown mess that will ruin your purification run, so err on a lower concentration.
Finally, the most obvious thing to do to clean up your prep is to do more washes with higher imidazole concentrations before you elute your protein. The most popular His-tag carries 6 histidine and proteins carrying these tags generally elute when the imidazole concentration reaches 100 to 180 mM, which means you can do washes at 30 mM, 50 mM, 70 mM and 90 mM imidazole before eluting your protein with 500 mM imidazole buffer. Each of these washes will remove some of the more stubborn proteins that don't interest you. If you see some of your protein already eluting at 90 mM or even 70 mM imidazole (this happens when the tag is partially hidden inside the protein after folding and only part of the His-tag is exposed, reducing its binding ability), then simply skip those step and add some extra washes below those concentrations. This is suitable for both beads and columns if you do this manually or with a peristaltic pump. However, if you have access to an FPLC system (e.g. Akta) you can run an imidazole gradient from 20 to 500 mM, which will generally produce a cleaner sampe and you see exactly at which point your protein elutes from the Ni-NTA column.
You could try to add some (0-10 mM) imidazole into the buffer you use when applying your protein to the column, use an imidazole gradient when eluting, or you could use TALON cobalt resins. Also you could just add another column (ion exchange and/or size exclusion). Of course you need to check if the additional bands are not HIS-tagged degradation products or protein multimers. If nothing works, you might considering changing the tag (e.g. GST). Best of success!
I would recommend just to include Wash 3 step with the buffer:
20 mM HEPES, 100 mM Imidazole, and 0.5M NaCl.
As Stefan mentioned, adding a small amount of imidazole to your lysis/binding buffer is recommended by most Ni-NTA column/beads manufacturers. The exact amount varies from protein to protein, but general 10-20 mM imidazole (even 30 mM) is tolerated by most proteins and it greatly helps to reduce non-specific binding.
Also, high NaCl concentrations help with this problem. You use 500 mM in your wash and elution buffers, but only 200 mM in your lysis/binding buffer. I always use 500 mM in all the buffers and, for some proteins that are highly charged (i.e. proteins with pI > 9) I even use 1 M NaCl.
Some of the contaminating proteins could actually be bound to your protein rather than to the Ni-NTA matrix. Adding a small amount of a reducing agent will help with this. For Ni-NTA I use a maximum of 3 mM betamercaptoethanol (BME) in my buffers, higher concentrations of BME or using DTT instead of BME will react with your Ni-NTA matrix and produce a brown mess that will ruin your purification run, so err on a lower concentration.
Finally, the most obvious thing to do to clean up your prep is to do more washes with higher imidazole concentrations before you elute your protein. The most popular His-tag carries 6 histidine and proteins carrying these tags generally elute when the imidazole concentration reaches 100 to 180 mM, which means you can do washes at 30 mM, 50 mM, 70 mM and 90 mM imidazole before eluting your protein with 500 mM imidazole buffer. Each of these washes will remove some of the more stubborn proteins that don't interest you. If you see some of your protein already eluting at 90 mM or even 70 mM imidazole (this happens when the tag is partially hidden inside the protein after folding and only part of the His-tag is exposed, reducing its binding ability), then simply skip those step and add some extra washes below those concentrations. This is suitable for both beads and columns if you do this manually or with a peristaltic pump. However, if you have access to an FPLC system (e.g. Akta) you can run an imidazole gradient from 20 to 500 mM, which will generally produce a cleaner sampe and you see exactly at which point your protein elutes from the Ni-NTA column.
I agree with adding imidazole into your lysis buffer. Also, we have been having a lot of success lately with a 1.0 - 1.5 M NaCl wash. Our buffers and protocols looks something like,
We use the 5 mL HisTrap columns from GE and the 25 mM Imidazole is what they recommend for eliminating non-specific interactions.
Good Luck!
If you have a band at ~75 kDa it's normally ArnA - I'd recommend adding glycerol (e.g. 15% v/v) to your buffers which can help your prep as it reduces hydrophobic interactions between proteins, even if it flows a bit slower (it will also help stabilise your protein of interest). Also as a warning, you will probably have another protein, SlyD, in your prep which is 26 kDa so will sit very near/on top of your protein on SDS-PAGE - you may be able to see a double band on high % Coomassie stained gels - if so, using 5 mM Bme in all your buffers can improve the situation a bit as it is a Cys rich chaperone. Both ArnA and SlyD are still His rich though so totally agree with other comments about Co resins/more washes+imidazole/extra purification step. You might also want to have a look at this DE3 strain with the main His contaminants fused to chitin binding domains, I haven't tried it myself but it seems good.
Good luck.
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3127686/
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