And after short time they became a legion (and expressed my protein!)

Oh yeah!

the magic of recombinant expression using the little bugs. Pre-culture, inoculate, induction and make kilos of the damn protein!

Today I want to share my optimized protocol for auto-induction medium for E.coli recombinant expression (T7 promoter system, of course!). I learned this protocol at an EMBO Course a the AFMB in Marseille and since then, and we’re talking about quite many years ago, I became a real fan of it, for a simple reason:

IT DAMN WORKS! 🙂

so, here is the recipe for this auto-induction medium. It is called ZYP5052, as it is the combination of ZY (or YT if you prefer), Phosphate/Sulfate buffer (NPS), and a mixture of glucose, lactose and glycerol, named 5052. The components must be prepared separately and mixed before inoculation.

Solution Name	Volume	Components	Amount
ZY	1 Liter	Tryptone Yeast Extract	10 grams 5 grams
1000xMg	100 ml	MgCl2	9.52 grams
20xNPS	1 Liter	(NH4)2SO4 KH2SO4 Na2HPO4	66 grams 136 grams 142 grams
50×5052	1 Liter	Glucose Lactose Glycerol	25 grams 100 grams 250 grams

Suggestion: the two solutions of NPS and 5052 will require a lot of time to dissolve properly, so I recommend to use pre-warmed water (heat it in a microwave) to prepare them. All this solutions, except the 1000xMg, must be autoclaved separately. The 1000xMg can be Filter Sterilized. The following quick reference will be of help when preparing the complete medium before the inculation:

Component	100 ml	250 ml	500 ml	1 Liter	5 Liters
ZY	92.8 ml	232 ml	464 ml	928 ml	4640 ml
1000xMg	0.1 ml	0.25 ml	0.5 ml	1 ml	5 ml
50×5052	2 ml	5 ml	10 ml	20 ml	100 ml
20xNPS	5 ml	12.5 ml	25 ml	50 ml	250 ml

It is of extreme importance that you mix the various components in the correct order, otherwise the local high concentration of Mg added to the NPS will result in precipitation and your medium will be cloudy (may still work, but don’t recommend it).

And now, here is my strategy for cell growth:

start an overnight culture of your favourite E.coli strain transformed with your expression plasmid. Use a volume corresponding to 1/50 of your final culture volume.

Inoculate your pre-culture in the ZYP5052 + antibiotics, shake vigorously (I use 200-250 RPM) the culture at 37 °C for 3 hours, then lower the temperature to 20 °C and let the culture continue until the next day.

After the overnight growth at 20 °C, harvest your cells. I experienced optical densities between 2 and 25, depending on the toxicity of the protein to be expressed, the E.coli strain, the growth condition (flask vs. bioreactor).

Good luck, and if you need a citation for the original method for E.coli auto-induction, here it is: Studier, F.W. (2005) Protein production by auto-induction in high-density shaking cultures Prot. Expr. Purif., 41, 207-234

Who to blame this time?

Every lab has its own stories about disasters or troubles due to lack of communication between the scientists. For many of us, the lab is our home, and our colleagues are like housemates. We have to trust them, we help each other, we talk, we make jokes, but we also keep an eye on what everybody else is doing, because we don’t want to run into problems (like dirty columns…) because of somebody’s distraction or laziness.

And when something is missing, or finished because the last person that took it did not order it promptly, the tragedy is around the corner. Obviously, nobody will be guilty, and actually wasting time trying to identify who did it will not solve the problem, it will just worsen the atmosphere. Moreover, your colleagues will most likely know that it’s you anyway.

Think about it next time you finish the last aliquot of a precious reagent, or when you put back a column after a purification, or while you go home without cleaning up a common space. Some simple and quick actions will avoidbig discussions and disappointment. And probably you’ll have more friends around you… 😉

Oh no, I lost my protein!

It’s funny. Every time I discuss about protein purification with someone I realize that there are extremely efficient ways to waste all your precious material in seconds, just because of distraction, lack of experience, or untrustable colleagues. And it seems that “becoming experienced” on the Akta means wasting bucketfuls of proteins from time to time.

So, I decided to prepare this “Top 5 reasons of failed protein purification using the Akta system” post and I hope that will help newbies to save their protein during their next purifications. Let’s start:

Unwanted air in the system that masks your protein peaks

Ok. Everybody knows that air and Akta are not compatible, but how many times a non-experienced user lets air inside the system? Sometimes people think that it’s just the column that should be kept free of air, but this is absolutely not true. An example: a 100 microliters loop left in “load” mode without syringe for a few seconds will introduce enough air in a MonoQ 5/50 column to generate annoying air spikes for the next 10 column volumes. Can you imagine the amount of air that can be introduced in a system just by changing a connection tube before or after your column, or replacing a loop?

Before your run, ensure you connected all your tubings and loops to the system. Bypass the column with a connector (the system should be “closed”) and flush the system with your buffer for at least 5-10 mls (should be enough to rinse all tubings from pumps to fraction collectors on most akta systems) with a high flow-rate (5-10 ml/min, unless you have special setups that require lower flow-rates, check your system!!!). The high flow-rate is required to remove occasional air bubbles trapped in the connectors or in the detector cell.

The sudden “after pumpwash” elution

One of the most horrible ways to lose your protein is see it coming out when you don’t expect it, and the Akta actually does not really help in avoiding it. The typical scenario is the following: IMAC, batch binding of a protein sample and manual packing of a column, you connect it to the Akta, your buffers A and B are on pumps A and B. Pumpwash, then start washing your column with buffer A, to remove non-specific contaminants before the elution with buffer B. Wait, that peak is too high, it’s really strange… anyway, let’s keep going, the UV signal is now flat, let’s do the elution with B… How come? there’s no peak! where’s my protein? Noooooooooo!!!!!! What happened?

Actually, I consider this a “bug” of the akta system: it depends on the dead volumes between pumps and valves. When you do a pumpwash, doesn’t matter which pump you will be using afterwards, the first 2-3 ml that will pass through the system (and through your column, if it’s connected) will be a mixture of your buffers A and B, which remains in the injection valve after the end of the pumpwash. If you are working with a 1 ml column (IMAC, MonoQ/S or something else), this tiny volume of concentrated elution buffer will elute all your sample, together with the damn contaminants, at the very beginning of your run. To avoid this unpleasant event, put your buffers on the Akta and wash the pumps before connecting your columns, or if your really need to swap your buffers during the run while your sample has already been applied to the column, disconnect (bypass) the column and flush the whole system for 5-10 ml after a pumpwash, and reconnect the column only after this step.

The mistery of the partial filling of the sample loop

I injected 500 micrograms of protein in 1 ml sample using a 1 ml loop, but I recovered only 300 micrograms… why?

This picture (taken from GE website) shows what happens to your sample when you apply it to a sample loop. Partial filling is the most efficient way to ensure 100% sample recovery using loops for injection.

Assuming that you really injected 500 micrograms of protein, that the sample did not stick to the plastic of the syringe, to the tubings or to the column beads, it may well be that you lost part of the sample because you sent it to the waste while filling the loop. Liquids actually do not migrate as straight lines in the akta tubes, and the more a tubing is narrow, the more the liquid will be retained by the inner surface of the tubing. That’s why partial filling of sample loops is the best way to ensure complete recovery of your sample during a run on the Akta. If you have a sample volume of 1 ml, use a 2 ml loop, and after applying your sample to the loop leave the system in inject mode for at least 3-5 times the volume of the loop (in the case of a 2 ml loop, use 10 ml). In many cases, you can leave the system in inject mode for the whole elution procedure (unless you need to use the sample valve for other reasons).

Loop bypass: from a syringe to the waste

Ok, we just discussed the partial filling of the sample loop, we are sure that there is no air in the system and the loop has been rinsed properly, by flushing it with a syringe from the injection port. To avoid the introduction of air in the loop, the wash syringe is still connected. So, the system is running, the column is connected, to remove my syringe from the injection port the system needs to be switched to inject mode, then I pick my sample, put the syringe in the injection port and push the plunger… The column is running, but nothing is coming out.

In load mode, the sample loop is connected to the injection port, which cannot be left open. In inject mode, the injection port is directly connected to the waste and the sample loop is bypassed.

Yes, it’s horrible to realize that your samples has never been injected in the column, because you forgot to switch to load before pushing the plunger of your sample syringe. In Inject mode, the injection port is connected directly to the waste, and the sample loop is completely bypassed. The correct procedure is the following: (1) while the system is in load, wash the sample loop using a syringe with a volume  capacity larger than the loop itself. (2) when it’s time to apply your sample, switch to inject, remove your wash syringe and put the sample syringe in the injection port, but do not push the plunger. (3) switch the system to load, and push the plunger of your sample syringe: your sample will go inside the loop. (4) switch the system again to inject to apply the sample to your column.

What if your colleague left a dirty column?

Your colleagues can be good friends, comrades, very helpul collaborators and will keep the spirit up during your daily life. However, you cannot trust everyone in everything, in particular when it regards doing cleanups or boring washing procedures. This means that, the more a column requires cleaning, the more you will find it dirty when you will really need it. And it will be horrible to see your protein on an SDS gel with additional bands of dirt after column processing (and trust me, it happens…)

So, if you are processing the sample of your life, do not trust your colleagues, do not trust your friends, do not trust your technicians. Wash your columns throughly before your run and make sure that they are properly clean when your sample will be applied.

Add tricks to the tricks!

This paper reports on the first crystal structure of a GPCR in complex with a G-protein. Amazing stuff. The “methods” section of the paper shows an incredible collection of tricks to stabilize, concentrate, crystallize and collect data on such a beast. Applause!

Faster and faster and faster…

Besides being the title of the new, very nice hit from Within Temptation, it’s also one of the standard skills that you have to develop in a competitive lab if you want to see your boss happy and famous.

Very often doing things faster means “quick and dirty”, which is not the case that I am going to describe. One of the most annoying procedures that I have to do on a daily basis is to stain/destain protein gels with coomassie blue or one of its friends. Methanol, stains everywhere, waiting for hours before seeing your faint bands, feeling the breath of your boss behind your shoulder that wants to know more than you if the protein of your dreams is being purified or not.

I use the microwave. So? nothing new, many people do. But I’ve seen several places where microwave staining is prohibited because of the boiling methanol, or because of the mess  that the staining/destaining procedure generates. But… I have a protocol that:

• minimizes the mess
• ensures the highest sensitivity
• will not kill you because of the vapors of methanol
• is damn FAST!!!!

so, here it is:

• after running your gel, put it in a glass beaker or even better in one of those glass containers used to make ice baths, filled with ~1 cm of deionized water
• microwave the gel for 1 minute, max power (which means, on my microwave, 1000W)
• wash out the water, then add 40 ml of Fermentas PageBlue solution. Cover your container with a paper towel (do not seal it), and microwave 3 times for 15 seconds at maximum power (1000W), gently shaking the container between the cycles. The blue stain should never boil. If you see that it starts boiling, reduce the length of the cycles.
• collect back the blue stain solution, rinse the gel with water (again, 1 cm in the container), and microwave for 30 seconds. Change the water again 1-2 times. it’s done!!!

The whole procedure takes less than 5 minutes and gives beautiful gels. The PageBlue solution is much more sensitive than standard coomassie R250 and can be re-used an incredible number of times. This makes this staining solution cheap despite the cost of a 1L bottle (I am staining 3-4 gels per week and I’ve been using the same 50ml tube of PageBlue since April).

I compared the staining results with Coomassie blue, Blue stain and PageBlue (without microwave), and I am proud to say that the sensitivity is very high (a 10-20 ng band is visible) and the background very low. It’s very important to wash the gel with water before staining to maximize the sensitivity. And it’s very clean! With a little piece of paper towel on the beaker, the solution will not spill around and your microwave will be as clean as new! And it’s not as toxic as standard Coomassie staining, because there’s no methanol!!!!

So, are you in a hurry with your SDS-gels? Give this protocol a try, and I’m sure it will not be the only time that you will use it!

Nanodrop: cannot imagine my daily experiments without it…

The NanoDrop ND1000 Spectrophotometer

It’s funny. Last week, when I published the post about the akta micro I thought that I should write something about another essential piece of hardware that we have in the lab (the Nanodrop), and just a couple of days ago the CCP4bb was flooded with messages discussing features and capabilities of this spectrophotometer. So, I think that it’s absolutely timely to describe why I think the NanoDrop is indispensable in a Structural Biology lab.

I can proudly say that in my previous lab in Pavia we pioneered the usage of NanoDrop when we purchased it in late 2005: our lab was the first Italian lab with such a spectrophotometer, and in the next years several other labs from Northern Italy purchased it after our recommendation. I think that I did not spend a single day in the lab since then without making at least one measurement at the NanoDrop.

How does it work?

It’s super easy! Every measurement requires 4 steps:

1. Check the system calibration using pure water (this step is required every time you access one of the measurement applications of the NanoDrop).
2. Blank the instrument with your buffer solution.
3. Make your measurement!
4. Remove your sample by wiping the pedestal and the lid with lint-free tissue (see below).

How long does it take?

It’s damn fast! A complete measurement does not take more than 2 minutes, including making the blank and cleaning-up after the measurement. It definitely takes MORE time to wash a quartz cuvette (without making a measurement). Moreover, as the nanodrop uses a Xenon lamp, no warming-up is required. The system is always ready for use.

Check samples in real-time

This is a short list of measurements that I perform routinely simply because the NanoDrop is so fast that I can do them while I’m doing other things. 

• Evaluation of sample concentration during concentration steps to see if the sample is doing well or if it is sticking to the membrane of the concentrator.
• Sample concentration before injection for analytical gel filtration
• Sample concentration immediately before crystallization
• DNA concentration (doing 20-30 minipreps per day, I would probably turn into a serial killer if I had to measure all my samples without a NanoDrop….)

Without the NanoDrop, I would probably skip most of the non-essential measurements. The result would be lower knowledge of the samples that I’m working with, and at the same more time spent to perform standard measurements with cuvettes.

Almost no protein consumption

Imagine a standard situation: the purification of your favourite protein is just finished, and you are working on a very challenging target, some of those nasty proteins that express in picograms per liter regardless the expression system. After months of struggle, you finally manage to have enough material to setup a couple of plates, and you want to measure the concentration of your sample before dispensing your precious 20 microliters with your favourite crystallization robot. With the nanodrop, in less than 5 minutes you get the concentration of your protein by using less than 1.5 microliters of sample, and if you are careful enough you can actually recover most of it after the measurement.

Closeup of the NanoDrop "cell". The sample is dispensed on the pedestal, then the measurement is performed. A magnet on the pedestal moves the lid to generate the 1 mm liquid column, where the first measurement is collected. Then, a second measurement with a path length of 0.2 mm is performed. (caption taken from the web, could not access the original source)

So, why many people describe issues about the reliability of NanoDrop measurements?

It’s the same old story: depending on your purpose, you may find the NanoDrop a great piece of equipment or a completely useless dustcatcher. You cannot use the NanoDrop to measure enzyme kinetics or to perform other analytical tasks, as well as you cannot rely on NanoDrop measurements if your concentrations are low. The short pathlength of the NanoDrop does not allow to perform reliable low-concentration measurements. In my experience, NanoDrop is excellent to measure protein samples in the 0.5 – 50 mg/ml concentration range (I’m talking about BSA-like proteins, with extinction coefficients around 1 M^-1cm^-1).

To get reliable results with the NanoDrop, the measurement should be performed rapidly, and the reproducibility should be checked by changing the sample, not by measuring the same drop 2-3 times. This will result in evaporation of the sample and subsequent increase in the sample concentration.

Another essential requirement if you want your NanoDrop to measure correctly is to clean it in the right way. Removal of the sample from the pedestal of the NanoDrop should be done immediately after sample measurement, by wiping from the back of the instrument to the front using a lint-free tissue. This last point is critical to ensure reproducible results over time. Normal paper tissues are not suitable to clean the spectrophotomer, as they will result in deposition of micro-fibers that will interfere with correct absorbtion measurements. We currently use the KimTech Delicate Task Wipers.

Maintenance?

The Xenon lamp will probably survive longer than many of the PostDocs and PhD students in the lab. The only essential check is the calibration setting. Every year, a check is recommended to see if the NanoDrop is measuring correctly. In case of large discrepancies, a technical intervention is required. However, if lint-free tissues are used and the pedestal is kept always clean and not washed with aggressive solutions, you’ll see that the instrument will maintain its calibration correct for many years.

Price?

People on the CCP4 described it as a costly instrument. In my opinion, it’s resonably cheap compared to a reliable multi-purpose UV/Vis spectrophotometer. Considering also that it is almost maintenance-free, its overall cost will be the initial purchase. But the increment in productivity and efficiency over time will pay for it, in short time.

NanoDrop it’s like it’s Hot!

This YouTube video is a cool summary of the features of the NanoDrop (thought was made by lab geeks like me, it’s actually sponsored by NanoDrop…). Anyway, enjoy:

CONECT: where Coot and PyMol collide

Coot is a fantastic program. That’s why most crystallographers use it during their daily tasks.

PyMol is also a fantastic program, with a very simple syntax and easy access to almost all its functions, it is my favorite publication-quality renderer.

I use to jump from coot to pymol and back several times when I prepare my pictures, and I believe that many of you do the same. There is one annoying issue that has been mentioned in the bulletin boards several times (see here and here for example) that is still waiting for a solution.

Let me describe it:

All the PDB files deposited in the Protein Data Bank that contain HET groups interacting with protein atoms, and/or contain disulfide bonds have at their end a section that contains CONECT records. These records strictly depend on the atom numbers involved in the record. Besides these records are an integral part of the PDB file format, in my experience they are hardly ever used fin protein crystallography for particular purposes. And here comes the issue: when you open a PDB file downloaded from the Protein Data Bank with Coot, the program will read the CONECT records, but will NEVER alter them. However, during most editing operations, the atom numbers contained in the file will be modified, and when the changes will be saved, the resulting file will contain a whole set of wrong CONECT records.

Is it a problem?

Well, if you re-open the file with Coot, not at all. But other programs like PyMol or UCSF Chimera consider the CONECT records as an integral part of the information that is read to display the object described in the PDB file.

After saving edits in Coot, a PDB file displayed in PyMol may show a crazy connectivity... This example shows PDB file 2XWB downloaded from the Protein Data Bank withn coot, superposition of chain J to chain A of PDB file 2XW9 (also downloaded from the PDB), modified file saved with Coot and opened subsequently in PyMol

The result?

Have a look at the picture here on the right. The connectivity is quite fancy, isn’t it?

How to solve it?

Here comes the easy part, just delete the CONECT records. But, as this operation is so easy, it becomes tedious if you have to do it manually many many times with a set of files under analysis. And as Coot at the moment does not alter the CONECT records at all (i.e., messes them up after coordinate edits), it would be great if these records were kicked out of the PDB files saved after a Coot editing session.

I’m sure it is in the “todo” list for one of the next Coot versions, I’m just impatient…

Why I like the Akta Micro

Last year we bought a new Akta system for protein purification. As we were looking for something to be coupled with a Multi Angle Laser Light

Akta Micro from GE Healthcare

Scattering system (MALLS), we were suggested by GE Healthcare representatives to buy an Akta Micro.

At the beginning we were really skeptic, because from the outside the machine is very similar to a standard Akta Purifier, but costs about 1.5 times its price. In the end we bought it (this GE reps can be very persuasive…) and after 1 year of intensive usage, I can say that it isa very good investment and I would like to share my impressions of this nice piece of equipment.

So, in practice, what are the differences?

Well, if you are looking for an add-on to your lab equipment, to perform analytical analysis of your protein samples (forget about doing initial sample polishing from lysates or media), then the Akta Micro is a good choice. The whole system setup is designed to minimize dead volumes, resulting in a super-high sensitivity compared to a standard Akta FPLC or Purifier 10. The three-wavelength UV detector (the cell path is 3 mm) allows to perform reliable analytical gel-filtrations with the amount of sample that you would normally use for an SDS-PAGE.

Does it work with any FPLC column?

No. It’s an analytical system and should not be used with preparative scale columns. Forget about low-pressure columns (HiPrep/HiTrap and similar, I refer to GE Healthcare columns only because that’s our setup at the moment), the pumps develop too much pressure and your columns may not survive a single run. The Micro system works great with analytical gel-filtration columns (PC 3.2 series, 5/150) and can also be used with semi-preparative columns (10/300). It does a great job also with IEX columns such as Mono/Mini beads.

Known problems?

The system develops VERY high pressures compared to a standard Akta system, which means that the risk of destroying an expensive column is just around the corner. It actually happened a couple of times in the lab with the PC 3.2 gel-filtration columns. The maximum flow-rate for these columns is very low (0.1 ml/min). By mistake (wasn’t me :-P), the flow-rate was set at 0.5 ml/min for less than a minute: the bed of the column packed of about 1 cm, leaving a gap between the top connection and the gel-filtration matrix. Horrible. More horrible the fact that these columns are sealed, which means yes, if it happens there’s no other option than buying a new column. And, as you know for sure, these columns are not cheap.

What about the coupling with a MALLS detector?

MiniDawn TREOS MALLS detector from Wyatt

We built our system by combining the Akta Micro with the MiniDawn Treos MALLS system from Wyatt coupled with a RID-10A dRI detector from Shimadzu (cheaper than the Wyatt dRI solution) and we are happy with our setup. The molecular weights calculated from the MALLS analysis are extremely accurate, and the chromatogram of MW distribution overlayed to the standard UV peaks gives nice ideas of the homogeneity and dispersity of the protein sample.

What about SEC columns for MALLS experiments?

We were suggested (from Wyatt representatives, actually) to use Wyatt SEC columns instead of GE analytical gel-filtration columns for GF-MALLS experiments. The reasons they told us were related to the fact that the silica-based columns from Wyatt minimize particle leakage that may interfere with MALLS detection during a gel-filtration experiment. In terms of resolution, my impression (at least with my samples) is that the two sets are comparable. However, there are some compatibility issues related to silica-based columns (a useful discussion from the CCP4 bulletin board summarizes them).

GE Healthcare Precision Column Holder

Another difference is that while Wyatt columns are “plug and play”, the analytical GE columns PC 3.2 require the connection through the Precision Column Holder, a sort of metal jacket to prevent damage due to high pressures. As we have both GE and Wyatt gel-filtration columns available, we decided to dedicate the Wyatt set for MALLS experiments as a default, whereas users that don’t use MALLS perform their analytical gel-filtrations using the GE set. Besides some sample-specific problems, my impression is that the GE and Wyatt columns are quite similar. I personally prefer the Wyatt columns because they allow higher flow-rates (0.5 ml/min), thus the equilibration and wash steps are much faster.

How often is the system used?

With approximately 10 users in the lab, and other 3 Akta’s available 24/7 for standard protein purifications, such a delicate and sensitive system may look like something that is used only every once in a while. Instead, the system if fully booked every day. At the moment the Akta Micro is used for about 90% of its time without MALLS connected, for small-scale analytical gel filtration of protein samples. The remaining 10% of the istrument time is used for MALLS analyses, in particular to check the quality of concentrated protein samples immediately before/after setting up of crystallization plates.

(another) salt crystal…

A horrible case happened few months ago: nice, birefringent crystals, they didn’t show up in absence of protein. However the X-ray shooting was pretty clear. It’s damn salt. Phosphate is a bad guy…

yet another salt crystal in my crystallization set-ups

Crystallization plates:
Corning 3550 Sitting-drop
Drop size (microliters):
0.15 protein + 0.15 reservoir
Dispensing technique:
Non-contact (Cartesian Honeybee)
Reservoir condition:
330 mM Ammonium Phosphate
33 mM Sodium Acetate pH 4.5
Protein Buffer condition:
25 mM HEPES/NaOH pH 7.8
50 mM NaCl
1 mM Nickel Sulfate
Protein Concentration:
8 mg/ml

Let ’em jump into your loop

Fishing crystals can be a painful operation, in particular if the crystals are very small. In this paper the authors found a very nice trick to make the crystals jump into the loop using sound waves… Impressive.