Difference between revisions of "Moore:Working with coli"
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Revision as of 13:50, 2 July 2008
--Smoore 21:43, 2 July 2008 (UTC)
E. coli is a Gram-negative bacterium classified as γ-proteobacteria. The closest to "wild-type" strains are probably a clinical isolates, but in the lab "wild-type" usually refers to a parental strain to which mutants are compared. Under ideal conditions (like in a fermenter), E. coli can replicate about every 20 minutes, but you are more likely to see a healthy strain divide about every 30-35 minutes in a well-aerated liquid culture at 37 °C. Most lab strains are considered harmless to human health (BL-1).
E. coli will grow at some rate anywhere from from ~15-45 °C until it has used all of an essential resource. After that, the bacteria enters a stationary phase wherein metabolism is greatly reduced and a series of physiological changes occur as the organism attempts to stay alive. Given enough time (a few days at room temp), a significant fraction of the bugs will die of starvation.
For most applications, the bacteria are growm in a medium containing yeast extract and tryptone as food sources (like LB broth). While this is an affordable and convenient way to make a lot of cells, care should be taken if these media are used during experiments. The pH and available carbon, nitrogen, and mineral resources are not controlled in these preparations so you can observe significant differences in behavior of the cultures from experiment to experiment. For controlled experiments, it is better to use a rich, defined medium that has all of the necessary components for growth. A buddy of mine and I set up an experiment to see how dense a culture would get if resources were not limiting. We put a healthy lab strain in a dialysis bag and dialyzed the culture agains fresh growth medium every day. After a week, the culture had stopped getting denser: when we measures the density of cells, the "OD" (described later) was ~85. The culture was still liquid though (a thick one) so the cells were not as dense as they are in a colony on a plate.
A Rant On Antibiotic Selection
A lot of words here, but important stuff to consider
Many times, you will have a need to maintain genetic elements that are linked to a gene that confers antibiotic resistance (for example a plasmid harboring a gene you want to express containing a "drug marker"). You should always keep in mind what the selection antibiotic is, how it inhibits bacterial growth, and what the resistance gene does to allow growth.
I have heard, many, many times, of people following protocols for protein expression where they are instructed to use "freshly-transformed cells". Think about it, how would the length of time a plasmid resides in a cell have any influence on the ability to induce a gene. In any given cell, the plasmid is as old as the chromosome, it doesn't "break" in the freezer stock to become un-indicible. This lab lore stems from the fact that plasmids can be lost in cultures if they are not selected for properly, especially if the cell doesn't like them.
A notorious bugger when it comes to this phenomenon is the use of ampicillin and β-lactamase for selection. Ampicillin inhibits a transpeptidase needed to make the bacterial cell wall. The β-lactamase enzyme irreveribly breaks the lactam ring in the antibiotic rendering it ineffective. For the resistance enzyme to work, it needs to be able to inactivate the drug near the site of cell wall synthesis (the periplasm in E. coli), so the enzyme is secreted. The problem arises because in the absence of complete inhibition of the transpeptidase (low drug concentrations), the cells will live and divide slowly. As the drug concentration drops further, the cells not making β-lactamase can begin to grow normally.
So let's take the folowing scenario: You start an overnight culture in regular LB broth of a T7-inducible gene on a plasmid conferring ampicillin resistance. You squirt in 100-150 μg/ml ampicillin and go home. When you come back in the morning, you take an aliquot of the culture, dilute it 1/100 in fresh medium with ampicillin and grow the culture to mid-log phase for protein expression. When you run a gel to see how much protein you made, you get hardly any. The strange thing is, when you first used the strain, it made tons of protein. You erroneously conclude that the difference is this strain wasn't "freshly transformed" with the plasmid.
What really happened was the β-lactamase secreted from cells containing the plasmid destroyed all of the antibiotic in the culture relatively fast. I can't remember the author, but I read a paper that reported this can happen in a matter of a couple of hours. Every now and then, some T7 polymerase got made in your expression strain because of leaky repression. These few molecules of polymerase made quite a bit of mRNA from your plasmid, the cells become burdoned and slow down. As the ampicillin concentration drops, cells not harboring an expression plasmid begin to grow quite well. If they are able to grow as well or better than cells containing the expression plasmid, they will represent a significant portion of the culture in the morning. When you dilute into fresh medium, the cells with plasmids as well as any liberated β-lactamase will immediately start chewing up the antibiotic. As with the overnight culture, the cells without plasmids rely on the enzyme produced by other cells. Since they don't have to replicate plasmids or deal with intermittent T7 expression, they gain ground so that by the time you induce the culture, a large fraction may not even have an expression plasmid.
Some tips for avoiding/diagnosing this problem:
(1) Avoid using β-lactamase and ampicillin to maintiain plasmids that might be toxic or burdon the cells.
(2) Harvest your overnight cells and resuspend in fresh medium to remove any free β-lactamase. I have not seen any data supporting this action, what I have read suggests that practically all of the β-lactamase is still cell-associated (periplasmic), but it makes people feel better to know they are trying.
(3) Add 0.1-0.2% glucose to any cultures/plates not intended for protein expression, especially the overnight (most protein expression systems respond to catabolite repression).
(4) Plate a dilution of cells at the time of induction and harvest to see how many still have plasmids (inferred by being ampicillin resistant). You may see a lot of amp resistant cells when you induce, they all stop dividing upon induction, and the empty cells keep growing so your harvested material is mostly cells that hadn't expressed a thing.
(5) Use more drug. I routinely use 200 μg/ml ampicillin.
Using Translation Inhibitors During Protein Expression
Some of the best selective antibiotics block protein synthesis. Most of the resistance genes inactivate the drug, some, like tetR, pump the drug out of the cell.
If you are using one of these guys to maintain a plasmid, you can cut back on the dose when it comes time to induce the cells for protein expression. Even though hey have a resistance enzyme, some drug will be free in the cell and can inhibit translation. You should be aware of what concentrations of antibiotic are required to prevent growth and realize that dose even lower than that will still significatnly retard growth of non-resistant strains.
Most E. coli strains are stored as "glycerol stocks" at -80 °C. To make these, liquid cultures are mixed with glycerol (10-20% final) and frozen. A scraping of the "freezer stock" is then used to seed fresh cultures as needed. In the absence of a cryo-protectant (like glycerol), freezing will rupture the cell membranes and kill the bug. In fact, freeze/thaw is one method to promote cell lysis for extracting protein. After new strains are generated, it is a good idea to make feezer stocks from relatively fresh cultures. If you need to wait for a few days (perhaps for sequencing data), keep the cultures at 4 °C.
E. coli is a pretty easy bug to kill. Most laboratories have defined protocols for sterilizing cultures/colonies before they are disposed of, so be sure to make sure your safety officer is happy with whatever you choose to do. Some of my favorite methods: bleach, autoclave, 75% ethanol, Wescodyne, and flame.
Measuring Culture Density
The simplest way to monitor a culture density is to measure the turbidity or "absorbance" (A) of a culture at a wavelength of light in the visible range. The reason I put absorbance in quotes is that it is a classic misnomer: the wavelngths of light chosen to measure cell density are deliberately chosen in ranges where absorbance is negligible. What you measure is the attenuance (D) of light as it passes through the sample before it reaches the detector. The attenuance is a combination of all of the light lost from absorbance, scattering, and instrument geometry. You won't be able to win over an editor when it comes to labeling your growth plots with D, just be aware that the "A600" measured in one spectrophotometer can be significantly different for the same cell density in another instrument. It is a good idea to measure the growth of a culture in the spectrophotometer you will be using so you are aware how the readings relate to the growth phase of the bacteria. You may go so far as to plate dilutions of the cells at different "absorbance" values to be able to convert your data to colony-forming units (c.f.u.).
Another common way of reporting density is with the term "optical density" (OD). Generally, this refers to the apparent absorbance of the sample in a 1 cm path cell. There is a really confusing tendancy of people to report amounts of material (cells, DNA, protein, etc.) as "ODs". When you see this, what is meant is "the amount of material that when diluted into 1 ml of solution will have an absorbance of 1 per cm". Why they don't use a concentration term is beyond me.
There is no reason to stick to using only 600 nm as the wavelength for measure culture density, any wavelength the scatters in proportion to the concentration of cells is fine. You may want to use 450 nm when the culture is very dilute, and 700 nm when it's thick. You can re-scale values measured at one wavelength to another value quite easily. Rather than fitting your data to a Mie scattering formula, you can make an empirical curve for one culture and use the relative values at different wavelengths to adjust values at other wavelengths.