Experiment 1: Basic Techniques:
Dilutions, Streaking and Spreading
Dilutions
The diluate is the liquid to be diluted and the diluent is what is used to dilute it. A dilution is defined by the following relationship:
volume of diluate = volume of diluate
volume of diluate + volume of diluent total volume
For example, if you add 9 ml of saline (diluent) to 1 ml of culture (diluate), this gives a 1/10 dilution:
1 ml of culture = 1 ml of culture = 1
1 ml of culture + 9 ml of saline 10 ml total volume 10
A 1/10 dilution is usually written as 10-1 (which is called the dilution factor) and is referred to as a 10-fold dilution. Note that a five-fold dilution is 2 x 10-1, whereas a 20-fold dilution is 5 x 10-2.
A single-step dilution is called a simple dilution. These are often used to make buffers and media. However, with bacteria and viruses one usually has to make a series of dilutions to achieve the desired overall dilution factor. There are several reasons for this:
1) The concentration of bacterial or viral suspensions is often above 109/ml.
2) It is often inconvenient or expensive if any tubes contain over 10 ml.
3) Volumes of less than 0.1 ml are difficult to measure accurately.
Therefore, most dilutions of greater than 100-fold are made serially. The thing to remember about serial dilutions is that the total dilution is equal to the product of the individual dilutions.
Problem: We need a 10-3 dilution of a culture
Solution: Make consecutive 10-1 and 10-2 dilutions, in either order.
Dilution Total
Step this step dilution
1. 0.1 ml culture + 9.9 ml diluent 10-2 10-2
2. 1.0 ml of above dilution + 9.0 ml diluent 10-1 10-3
OR
1. 1.0 ml culture + 9.0 ml diluent 10-1 10-1
2. 0.1 ml of above dilution + 9.9 ml diluent 10-2 10-3
Culture Dilutions
To count the cells in a culture, the culture is diluted and a sample from the final dilution is plated onto agar. After overnight growth, the colonies are counted. If you know what dilutions were made, the volume plated and the colony count, you can calculate the concentration of cells in the original culture.
Usually you know the number of colonies on the plate. From this you can calculate the concentration of cells in the final dilution tube (i.e. the one used when plating out a sample).
If 1.0 ml of the dilution was plated out the plate count would be equal to the number of cells/ml in the dilution tube. In practice the volume plated out is usually 0.1 ml. [It would take much too long for the plate to dry if 1.0 ml was used.] When a volume other than 1.0 ml is plated, divide the number of colonies on the plate by the amount plated to get cells/ml.
Example: You plate out 0.2 ml from the final dilution tube and the colony count is 130.
What is the cell concentration in the final dilution tube?
Solution: The concentration in the final dilution tube = 130 cells = 650 cells/ml
0.2 ml
To calculate the concentration of cells in the original culture, you need to take into account the dilutions made before the final dilution tube.
The dilution factor gives you the fraction of cells in the final dilution tube. For example, if you made a 100-fold dilution (dilution factor = 10-2) from tube A into tube B, the concentration of cells in tube B equals the concentration in tube A divided by 100 (or x 10-2).
Initial concentration x dilution factor = final concentration
By rearranging the above equation you get:
Initial concentration = final concentration
dilution factor
Example: You make a 10-6 dilution from your original culture to get the sample you plated out in the example above. You know that the final dilution tube has 650 cells/ml.
Original concentration = (650 cells/ml) ÷ (10-6) = 6.5 x 108 cells/ml
To get accurate results you must thoroughly mix each dilution. You must use a clean pipette or a fresh pipetman tip for each dilution. To measure volumes accurately never use a pipette that has a capacity of more than ten times the volume to be pipetted. Make sure you don't transfer droplets clinging to the outside of the pipette or tip.
Sometimes your lab directions might say something like "plate a 10-4 dilution." Unless otherwise instructed, this would mean to plate 0.1 ml of a 10-4 dilution.
Quite often in the lab you want to dilute a concentrated solution to an exact concentration. This type of dilution can be calculated by the following equation:
volume A x concentration A = volume B x concentration B
Problem: You have a bottle of 95% ethanol and some distilled water. How much ethanol and how much water do you mix to make 100 ml of 70% ethanol?
Solution: (x ml) (95%) = (100 ml) (70%) therefore x = 7000 ÷ 95 = 73.7
Therefore mix 73.7 ml of 95% ethanol with 26.3 ml of distilled water.
Streaking for Single Colonies
Single colonies of a bacterial strain growing on agar almost always arise from a single cell. In most cases it is of fundamental importance for a geneticist to begin work with as homogeneous a population as possible. Therefore, you must be able to isolate single colonies.
As shown below, cells from a plate, stab or liquid culture are first applied in a streak using a sterilized loop (Step 1). If you are applying liquid, let it dry before the next step. Resterilize the loop, cool it (by sticking it into an unused portion of the plate), and cross-streak as shown (Step 2). Resterilize the loop once more, cool it, and cross-streak again, taking care to only touch the first cross-streak, not the original streak (Step 3). Repeat once more (Step 4).
If you wish to use a previously learned method, please do so. If you are familiar with this technique you should divide the plate in two down the middle and do two separate isolations on a single plate.
Spreading Cells on Agar
Spreading is a technique used to get separate colonies from a culture. The culture is first serially diluted. Then portions from suitable dilutions are spread over the surface of an agar plate using a sterile bent glass rod.
Sterilization: The spreader is sterilized by immersion in 95% ethanol. Just before use it is passed briefly through a flame to set the ethanol alight.
CAUTION: Do not hold the glass rod in the flame - you only want to burn off the alcohol – this leaves the spreader cool. The spreader is sterilized by the alcohol NOT by the flame. If the glass becomes too hot you will kill the bacteria you are trying to spread.
CAUTION: Make sure that no flaming ethanol drops into the beaker of ethanol.
The spreader is then used to distribute the liquid evenly over the surface of the plate. Use one hand to hold the glass spreader and the other hand to turn the plate as you spread.
Experimental Method: Dilutions, Streaking and Spreading
Materials per class:
4 L-broth plates per student
(L-Broth contains, per liter, 10g tryptone, 5g yeast extract & 5g NaCl)
Sterile saline (0.85% NaCl)
10 large sterile test tubes per student
Inoculating loops
Spreaders (i.e., bent glass rods)
Sterile 1 ml and 10 ml pipettes
95% ethanol
Glass Petri dishes (to hold the ethanol)
1 overnight culture of E. coli per lab bench
Pipettemen to measure 0.1 ml (= 100ml) volumes
Sterile tips for pipettemen
DAY 1 - Lab Period:
Safety Tip: When you insert a pipette into a pipette pump hold the pipette at the end closest to the pump. Turn the pipette gently as you insert it. If you hold it at the far end you are likely to break the pipette and you may stick the broken end into your hand and get blood all over the lab benches.
Assume the overnight culture has between 1 to 5 x 109 cells/ml. Make 10-fold dilutions such that spreading 0.1 ml of one of them should give you between 20 and 300 colonies. Bracket this dilution with one that should give you 10-fold more and one that should give you 10-fold less. Record the dilutions used. Spread 0.1 ml of each of these dilutions on three of the L broth plates. When the plates are dry, invert and incubate at 37˚C.
Divide your last plate into 2 equal sectors. In each sector do a separate isolation by streaking.
DAY 2 - Unscheduled Lab:
Count colonies and record the results. Calculate how many cells/ml were in the original overnight culture and post your calculations and results on the board in lab. For those of you who have never counted colonies or don't have time, refrigerate your plates in the cold room for counting in the next lab period. Do not leave your plates in the incubator as the colonies will overgrow. Refrigerate your colony isolation plates at this time also; show them to one of your lab instructors during the next lab period.
Experiment 2: Phenotypes and Genotypes
Scientists need to be able to communicate precise information. Two types of information that geneticists are concerned with are the genotype (genetic makeup) and the phenotype (observable characteristics) of an organism. In this exercise, you will first be introduced to the appropriate nomenclature for Escherichia coli and then you will determine if the reported genotype of a strain is correct. The rules for the genetic nomenclature for E. coli and for Salmonella typhimurium were first set down in 1966. These have subsequently been expanded to take into account genetic manipulations that were unknown in the past.
Genotype. Three letter italicized symbols are used to describe genetic loci, e.g. arg and lac. By convention, you underline letters if you can't make them in italics, e.g. arg and lac. When necessary, the three letter symbol is followed by a single italicized capital letter to distinguish related genes. The genes argA and argB encode different enzymes involved in arginine biosynthesis. Allele numbers can also be added to distinguish between different mutations in the same gene. Therefore, lacZ213 and lacZ15 designate two different mutations of the gene encoding b-galactosidase. The plus sign is used to designate the wild-type allele, e.g. lacZ+ is the wild-type state of the lacZ gene. When writing out a strain's genotype, one usually lists only the known mutations, i.e. wild-type alleles are not given. Therefore, the mutant allele is not given a minus sign. The designation argA is sufficient to indicate a mutant allele; argA- is considered overkill.
Other nomenclature you should know. The symbol ∆ indicates a deletion of a gene or region, e.g. ∆ (argF-lac)U169 is a particular mutation in which part of the genome from argF to the lac operon has been deleted. The letter f indicates a gene fusion. E.g. f(adhE-lacZ) is a fusion of the regulatory region of adhE to the structural gene for beta-galactosidase. The symbol :: indicates an insertion, e.g. lacZ::Tn10 indicates an insertion of the transposable element Tn10 into the lacZ gene.
Phenotype. Phenotypes are written descriptions or abbreviations that sometimes resemble the symbol for a genotype, e.g. Arg- indicates that the strain cannot synthesize arginine and thus requires arginine for growth. The genotype could be argD or argA or be a mutation in any other of the many genes needed for arginine biosynthesis. Note that in order to understand a phenotype (i.e. the growth characteristics of a strain) you need to know something of bacterial metabolism. Arg- means the strain requires arginine. In contrast, Lac- means the strain cannot use lactose. You must also understand something of genetic regulation, e.g. most argR mutants would not have an Arg- phenotype. (Would you expect a lacI mutant to be Lac-?)
Checking a strain's markers. A bacterial geneticist usually employs strains that already carry some mutations. This greatly facilitates many genetic manipulations. When you are given such a strain, the first thing you do is streak it out to isolate single colonies. Then, you pick a colony and grow an overnight culture in an appropriate medium such as L broth. It is essential that you then check the strain's markers so that you are sure you will be working with the correct strain. A control for contaminated medium is done by incubating an uninoculated tube overnight.
In this exercise, we will check some of the markers of strain JK270. It is very unusual to check all of a strain's markers. You must check the ones that are important in any experiments you wish to do with the strain, and at the same time you usually check a few other markers that are easy to work with. It is important to also use one or more control strains. This is because some of the tests are negative tests. Strain JK270 is supposed to have a his mutation. However, if we plate it out on medium without histidine and it doesn't grow, does this necessarily mean it is His-? Perhaps the plate has no glucose or you have inadvertently added an antibiotic. You can control for this in several ways, one of which is to also use a strain that should grow on the plate. We will use as a control strain JK114, which is wild-type for all of the markers we are checking.
The genotype of JK270 is: (F-) galK, his, lacY, lysA, metB, mtl, proA, pyrE, rpsL, xyl thi
The characteristics of these mutations are listed below. (These are the abbreviations used by all bacterial geneticists.)
Genotype Phenotype Description of trait or enzyme affected
F- recipient does not contain the F plasmid
galK Gal- galactokinase
his His- mutation in one or more genes of his operon
lacY Lac- b-galactoside permease
lysA Lys- diaminopimelate decarboxylase
metB Met- cystathionine-g-synthase
mtl Mtl- mutation in a gene for mannitol utilization
proA Pro- g-glutamyl phosphate reductase
pyrE Ura- orotate phosphoribosyltransferase
rpsL StrR ribosomal protein S12, streptomycin resistance
thi Thi- requirement for thiamine (vitamin B1)
xyl Xyl- mutation in a gene for xylose utilization
It is obviously necessary to know the standard three-letter abbreviations for the amino acids as well as those for many common sugars and the three letter (as well as two letter) abbreviations for some antibiotics, e.g. Str for streptomycin. It is a good idea to begin familiarizing yourself with the E. coli genetic map. Once one knows the presumed phenotypes, one can design tests for the markers. The nutritional requirements of auxotrophs are easy. Make a glucose minimal medium plate with all of the requirements, and then a series of similar plates with each one missing a different, single requirement. Plates are used rather than liquid cultures because accidental contamination is easily detected on plates and can usually be ignored. Antibiotic resistance markers are also easy to score. The appropriate antibiotic is simply added to a rich medium plate.
The ability to use different sugars as a carbon source can be tested in two different ways. The first is to make a minimal medium plate with the carbon source you are testing plus any other growth requirements (usually amino acids or vitamins). You cannot use a rich medium like L broth with the sugar you wish in place of glucose because E. coli grows fine using different components of tryptone and yeast extract (both present in rich medium) as carbon sources. Fortunately, E. coli cannot use most amino acids as its sole source of carbon and energy. In addition, the concentrations of amino acids added as supplements are much lower than those necessary for visible growth using the amino acid as a carbon source. The second method is to use fermentation indicator plates. The sugar being tested on such plates must be one that E. coli ferments to produce acidic products; therefore, this method cannot be used for all carbon sources. We will use this method to test for use of lactose, galactose and xylose. These sugars will be added one at a time to MacConkey base medium which contains neutral red as pH indicator. Fermenting colonies excrete acidic by-products and will appear red. Nonfermenting colonies look white or colorless. Actually, MacConkey plates are selective as well as differential. The medium contains bile salts and crystal violet that kill Gram positive bacteria and certain E. coli mutants that have defective cell walls.
For a complete list of E. coli genes and other genetic elements see the series of chapters in Volume 2, Section VI of Neidhardt F.C., et al. (Eds.), 1996, Escherichia coli and Salmonella Cellular and Molecular Biology. ASM. Washington, D.C. 2nd edition.
Experimental Method: Phenotypes and Genotypes
Materials per pair:
Overnight cultures of JK114, JK270 and an unknown strain
Sterile tube for making dilution
Sterile saline (0.85% NaCl)
Inoculating loops
Sterile 1 ml and 10 ml pipettes
Pipettemen to measure 0.1 ml (= 100ml) volumes
Sterile tips for pipettemen
One each of the following plates:
L broth
L broth plus 200 µg/ml streptomycin
MacConkey lactose (1% wt/vol)
MacConkey galactose
MacConkey xylose
[Note: MacConkey agar mix contains lactose. To use other sugars, use MacConkey BASE agar mix which has no sugar added. Then add 1% wt/vol of the sugar.]
#1 = M9 glucose vitamin B1 with His, Lys, Met, Pro, Ura
#2 = M9 glucose vitamin B1 with His, Lys, Met, Pro, -
#3 = M9 glucose vitamin B1 with His, Lys, Met, - , Ura
#4 = M9 glucose vitamin B1 with His, Lys, - , Pro, Ura
#5 = M9 glucose vitamin B1 with - , Lys, Met, Pro, Ura
[Note: Glucose is used at 0.4% (must be autoclaved separately from M9 salts). Amino acids are used at 50 µg/ml final concentration, uracil at 10 µg/ml and vitamin B1 at 5 µg/ml.]
M9 Minimal Salts Medium (Miller, 1972) in g/L
Na2HPO4............................................................................... 6.0
KH2PO4 ............................................................................... 3.0
NaCl....................................................................................... 0.5
NH4Cl.................................................................................... 1.0
After sterilization, MgSO4 and CaCl2 are added to 1 mM and 0.1 mM respectively.
DAY 3 - Lab Period:
Divide each plate into three. Label where each of the three strains will be inoculated. Also mark the plates on the bottom with your name.
Make a ten-fold dilution of each of the overnight cultures into saline. This is necessary to reduce the carryover of unused nutrients in the L broth. Using a sterile loop, put a small drop of each strain in its designated place. It is important that the drops you add do not run together. As soon as the plates are dry, invert and incubate at 37˚C.
DAY 4 - Unscheduled Lab:
Check all of your plates and record the results. Are the results for JK270 what you would predict? What is the phenotype of your unknown strain? Using the reference, predict possible genotypes.
Experiment 3: Growth Rate & Cell Composition
Bacteria show wide variations in growth rate in response to changes in their environment. They may exhibit 20-fold or greater changes in growth rate depending on incubation temperature or medium composition. One general observation is that the number of ribosomes increases with the increased growth rate. This is because more ribosomes are necessary for the higher rate of protein synthesis required at high growth rates. Since ribosomes consist of approximately 70% RNA and 30% protein by mass the relative cellular content of RNA drastically increases. At elevated growth rates, the RNA/protein ratio therefore increases. In addition, individual cells grown at higher rates are larger, and have more RNA, DNA, and protein per cell.
In this experiment you will measure the amounts of RNA and protein at two different growth rates. Two cultures of E. coli growing in different media will be used to achieve different growth rates. Cultures in lag phase and early stationary phase contain a mixture of cells metabolizing at differing rates. For a meaningful comparison of cells grown at different rates, we need homogeneous cultures in which all the cells are in the same metabolic state. Such a situation may be achieved by subculturing the cells through several rounds of exponential growth, and then analyzing the cells while in mid exponential phase.
Overall Macromolecular Composition of an Average E. coli Cell
|
Macromolecule |
% Dry weight |
Weight / cell femtograms |
Molecular weight |
Number per cell |
Different Kinds
|
|
|
Protein |
55.0 |
155.0 |
4.0 x 104 |
2,360,000 |
1050 |
|
|
RNA |
20.5 |
59.0 |
|
|
|
|
|
23 S rRNA |
|
31.0 |
1.0 x 106 |
18,700 |
1 |
|
|
16 S rRNA |
|
16.0 |
5.0 x 105 |
18,700 |
1 |
|
|
5 S rRNA |
|
1.0 |
3.9 x 104 |
18,700 |
1 |
|
|
transfer |
|
8.6 |
2.5 x 104 |
205,000 |
60 |
|
|
messenger |
|
2.4 |
1.0 x 109 |
1,380 |
400 |
|
|
DNA |
3.1 |
9.0 |
2.5 x 109 |
2 |
1 |
|
|
Lipid |
9.1 |
26.0 |
705 |
22,000,000 |
4 |
|
|
Lipopolysaccharide |
3.4 |
10.0 |
4346 |
1,200,000 |
1 |
|
|
Peptidoglycan |
2.5 |
7.0 |
(904) |
1 |
1 |
|
|
Glycogen |
2.5 |
7.0 |
1.0 x 106 |
4,360 |
1 |
|
|
Total macromolecules Soluble metabolites |
96.1 2.9 |
273.0 8.0 |
|
|
|
|
|
Inorganic ions |
1.0 |
3.0 |
|
|
|
|
|
Total dry weight |
100.0 |
284.0 |
|
|||
|
Water at 70% of cell |
|
650.0 |
|
|||
|
|
|
_______________ |
|
|||
|
Total weight of one cell |
|
924.0 femtograms |
|
|||
Remember: 10-3 gram = milligram 10-6 gram = microgram 10-9 gram = nanogram 10-12 gram = picogram 10-15 gram = femtogram
KLETT COLORIMETER OPERATING PROCEDURE
Setting Up The Machine:
(Should be done by TAs; Leave the Zero Set knob alone unless there are problems!!)
1. Zero the Pointer with the Zero Set knob
3. Make sure Photocell Switch is OFF
5. Turn on Lamp Switch and allow to warm up for 15 minutes
Zeroing On The Blank:
2. Insert tube with blank into instrument
4. Turn Scale Knob so scale dial reads ZERO
6. Turn on Photocell Switch
7. Adjust the Pointer back to Centre Line using the Zero Adjust knob
To Take A Reading:
8. Turn OFF Photocell Switch before removing tube
9. Remove blank (or previous sample tube); Insert sample tube
10. Turn on Photocell Switch
11. Use the Scale Knob to bring Pointer back to Centre Line. Record the Scale Reading.
[Repeat the Scale Knob adjustment twice more and average the three readings]
12. Turn OFF Photocell Switch
Experimental Method: Growth Rate & Cell Composition
DAY 5 - LAB PERIOD:
Materials
Klett colorimeter with green filter (or Spectronic)
[NOTE: This experiment may be switched to using the Spectronic instead of the Klett colorimeter. If so you will be given modified instructions during the lab.]
Exponential cultures of E. coli grown in rich and minimal medium. Strain DC500 is used because it grows relatively fast. It is more or less wild type and has no growth requirements.
Minimal medium = M9 salts plus glucose (0.4%)
M9 Minimal Salts Medium (Miller, 1972) in g/L
Na2HPO4............................................................................... 6.0
KH2PO4 .............................................................................. 3.0
NaCl....................................................................................... 0.5
NH4Cl.................................................................................... 1.0
After sterilization, MgSO4 and CaCl2 are added to 1 mM and 0.1 mM respectively.
Rich medium (L Broth plus glucose)
Sorvall high-speed centrifuge with rotor
50 ml centrifuge tubes (2 per pair)
Ice bath
Sterile Side arm flasks with sponge bungs (2 per pair)
Sterile Pipettes (1, 5, and 10 ml)
Pipettemen to measure 0.1 ml (= 100ml) volumes
Sterile tips for pipettemen
Vortex mixer
Sterile saline (0.85%)
Sterile 18 mm tubes for dilutions
Glass spreaders, glass petri dishes and alcohol
L-broth agar plates (make sure these are dried properly!)
Growth Curve:
Each pair of students will grow one rich and one minimal culture in side arm flasks. The volume of each culture will be 25 ml. The culture density will be followed till late exponential phase using the Klett colorimeter. When the culture has reached 100 Klett units you will carry out a dilution and plating to estimate the viable count, and then centrifuge the rest of the culture.
1) Aseptically transfer 24.0 ml of rich broth and 23.0 ml of minimal medium into two separate side arm flasks.
2) Inoculate the rich broth flask with 1.0 ml of E. coli grown in rich broth and inoculate the minimal medium with 2.0 ml of E. coli grown in minimal medium, so making the total culture volume 25 ml in each case.
Note: If the starter cultures are of low density you may need to add more inoculum and less medium. If this happens your TA will inform you of the proper volume to use.
3) Read the absorbance of each culture immediately after inoculation in the Klett meter. Use the corresponding medium without cells for a blank. (TA will provide blanks). Your initial reading should be 10-20 KU.
4) Place the flasks in a shaking water bath set at 37°C and speed 6. Read the absorbance every 30 minutes (for the rich culture) or 45 min (for the minimal culture) until the culture reaches >100 Klett units (KU).
5) Once the culture is over 100 Klett units, remove 0.1 ml. Dilute this sample to obtain between 300 and 3000 cells per ml. Then plate out 0.1 ml of this dilution on L-broth plates for viable cell counts. You should also plate out the dilution above and below this one. (NOTE: to calculate the proper dilution assume that 100 KU = 5 x 108 cells/ml approximately).
6) Pour remaining cells into a sterile 50 ml centrifuge tube. Place in ice bath to stop further growth until the centrifuge is ready. Label the tubes with a felt marker NOT tape.
7) Sediment cells for 10 minutes at 10,000 rpm at 4°C in the centrifuge.
8) Resuspend cells in 10 ml of cold saline (on ice). Recentrifuge for 10 min at 10,000 rpm.
9) Pour off supernatant and place tubes with pelleted cells in freezer until next lab period.
10) Incubate your viable cell count plates overnight at 37°C. Come in the next day and store these in the cold room. Remember to collect and count these in the next lab period.
DAY 6 - Unscheduled Lab: Count the colonies on the viable count plates.
DAY 7 - LAB PERIOD: PROTEIN & RNA ASSAYS
Materials:
Distilled water
Klett meter with a 660 nm red filter
Klett tubes (10 per group)
Flask (100 ml)
Boiling water bath
Pipettes
Test tubes with plastic screw caps (24 per group)
Lowry protein assay reagents
Orcinol RNA assay reagents
Standard solution of adenosine (for RNA assay)
Standard solution of bovine serum albumin (BSA)
Prepare Cell Pellets for Assay
Resuspend the frozen cell pellets in 5 ml of distilled water. Remember when you come to do your calculations that your suspension of 5 ml is equivalent to 25 ml of original culture (less the 0.1 ml sample you removed - but you can ignore this as it is so small). Assay this cell suspension for RNA and protein. The assays used here are colorimetric, i.e. the intensity of the color produced is directly proportional to the amount of RNA or protein. Start the RNA assay first as it takes longer. You can set up the protein assay while the RNA assay is incubating.
A) Lowry Assay for Protein
Although there are many different proteins in any given cell, they all have one thing in common - the peptide bond which links the different amino acids together. Most protein assays actually test for the presence of peptide bonds. Every protein has, on average, the same number of peptide bonds per unit weight. The Lowry assay involves two reactions:
a) the reaction of peptide bonds with alkaline copper and
b) the reduction of phosphomolybdate by tyrosine and tryptophan residues.
Assay Materials
100 mM NaOH
Protein standard solution: 500 mg/ml of BSA (Bovine Serum Albumin)
Lowry A: 2% Na2CO3 in 100 mM NaOH
Lowry B: a 1:1 mixture of 1% CuSO4 and 2% Sodium tartrate, mixed fresh before use.
Lowry D: Folin-Ciocalteau phenol reagent diluted 1:1 with H2O just before use.
Assay Protocol
1) Make a 1 in 5 dilution of each cell suspension in distilled water. Add 2 ml of 100 mM NaOH to 1 ml of the diluted cell suspension. Use a tube with a plastic screw cap. Remember, for your calculations that this has diluted your cells 1 in 5 x 3, i.e. 1 in 15.
2) Place the mixture in a boiling water bath for 10 minutes. This destroys the cell wall. Use the solubilized cells as your sample. The volumes given in the tables below leave half your sample in case you need to repeat the assay. If you do so make a new blank tube.
3) Set up a standard curve in five tubes:
Tube Water (ml) BSA (ml) BSA (mg)
1 (blank) 1.0 0.0 0
2 0.9 0.1 50
3 0.8 0.2 100
4 0.6 0.4 200
5 0.2 0.8 400
4) Set up your experimental assay tubes:
Tube Water (ml) Solubilized Cells (ml)
6 0.5 0.5 minimal
7 0.0 1.0 minimal
8 0.5 0.5 rich
9 0.0 1.0 rich
5) Mix 50 ml Lowry A with 1 ml Lowry B in a flask. This gives Lowry C.
6) Add 5 ml of Lowry C to each tube and vortex. Stand 10 minutes at room temperature.
7) Add 0.5 ml Lowry D and vortex. Stand for 30 minutes.
8) Zero the Klett meter (with the 660 nm red filter in place) using Tube 1 as the blank.
9) Read the absorbance of the assay tubes.
B) RNA-Orcinol Assay
The orcinol assay measures the concentration of ribose (deoxyribose gives about 5% the reaction of ribose; since the amount of DNA in a cell is much less than that of RNA, its impact on the assay will be negligible). When heated in concentrated acid, pentoses form furfural. This reacts with the orcinol in the presence of FeCl3 to give a green color. This reaction is restricted to free pentoses. Acid treatment releases the ribose from purine nucleosides, but not from pyrimidine nucleosides. Hence only half of the RNA reacts. Since adenosine is used as standard and will all react, the actual amount of RNA is twice what the assay indicates.
Assay Materials
10% TCA (trichloroacetic acid)
RNA standard: 100 µg/ml adenosine (equivalent to 200 mg/ml RNA)
Orcinol reagent : 1 gm orcinol in 5 ml of 95% EtOH mixed with 100 ml of
concentrated HCl containing 0.1% FeCl3. Make this solution fresh daily.
Assay Protocol
1) Add 2 ml of water and 3 ml of 10% TCA to 1 ml of cell suspension in a test tube with a screw cap. Remember, for your calculations that this has diluted your cells 1 in 6.
2) Place in boiling water bath for 20 minutes and allow to cool for 10 -15 minutes.
3) Centrifuge for 10 minutes at 3/4 speed in a clinical centrifuge. Be sure there is a rubber pad beneath the tubes. Pour the supernatant into a fresh tube. The volumes given in the tables below leave half your sample in case you need to repeat the assay. If you do so make a new blank tube.
3) Set up a standard curve in five tubes:
Tube Water (ml) Adenosine (ml) RNA equivalent (mg)
1 (blank) 2.0 0.0 0 x 2 = 0
2 1.5 0.5 50 x 2 = 100
3 1.0 1.0 100 x 2 = 200
4 0.5 1.5 150 x 2 = 300
5 0.0 2.0 200 x 2 = 400
4) Set up your experimental assay tubes:
Tube Water (ml) Solubilized Cells (ml)
6 1.0 1.0 minimal
7 0.0 2.0 minimal
8 1.0 1.0 rich
9 0.0 2.0 rich
6) Add 1.5 ml orcinol reagent to each tube and place in boiling water bath for 30 minutes.
7) Allow the samples to cool, then add 4.0 ml water.
8) Blank the Klett meter (with red filter in place) using Tube 1 (the blank).
9) Read the absorbance of the assay tubes.
Lab Write-Up
1) Plot the KU vs. time for each culture on 2-cycle semi-log paper. Read the mean generation time, g, from this plot and hence calculate the specific growth rate, m, for each culture. You will have to find the equation to derive m from g on your own. The specific growth rate, m, is also sometimes known as the growth rate constant, k.
NOTE: Plot data directly on log paper. Do not calculate logarithms yourself – the log scale paper does this for you!
NOTE: Do NOT calculate the mean generation time – get it from the graph. (WHY??)
2) Calculate the viable count for each culture.
3) Plot the standard curves for each chemical assay on linear graph paper. Calculate the amounts of RNA or protein in each of your experimental samples.
4) Calculate the total amount of RNA and protein per 1.0 ml of the ORIGINAL culture. Remember you resuspended the cells in a different volume after harvesting them. You also diluted them during acid / alkali treatment just before each assay. Calculate the RNA/protein ratio for each culture. Comment on these ratios.
5) Calculate the number of cells per Klett unit for the two cultures. Calculate the amount of protein per cell for each culture. What does this tell you about the relative cell size in the two cultures?
Experiment 4: The lac Operon and b-Galactosidase
The Lactose Operon
The lactose operon encodes three proteins, LacZ - b-galactosidase, LacY - the lactose permease, and LacA - lactose transacetylase. The operon is repressed by the LacI repressor whose gene is close to, but not part of the lac operon. Presence of inducer inactivates the repressor. Lactose itself is not an inducer and only indirectly induces the lac operon after a small amount has been isomerized to allolactose. This is an isomer of lactose, generated in a side reaction by the low basal levels of b-galactosidase which are found before induction. In the laboratory, IPTG (isopropyl-thio-b-D-galactoside) is often used as inducer. IPTG is not metabolised and is of no use to the cell - it is a gratuitous inducer.
Lactose enters the cell via the inducible lactose permease. Lactose entry is coupled to the proton motive force. The respiratory chain generates the PMF by pumping H+ ions out from the cell. The lactose permease operates by proton symport. An H+ ion, re-entering the cell provides the energy to move the lactose across the membrane.
Catabolite Repression
Presence of a favored carbon source such as glucose prevents use of less favored substrates such as lactose. Catabolite repression depends largely on the intracellular level of cyclic AMP. Cyclic AMP is bound by Catabolite Activator Protein (CAP) also known as cyclic AMP Receptor Protein (CRP). The level of CRP is constant. Transcription of catabolite sensitive operons such as the lac operon requires binding of CRP-cAMP complex to the promoter region. This allows RNA polymerase to bind to and transcribe the operon.
Carbon Source Cyclic AMP Level Target Gene
GOOD LOW OFF
BAD HIGH ON
The regulation of cyclic AMP levels is due mostly to changes in activity of adenylate cyclase which catalyses the conversion of ATP to cyclic AMP plus inorganic pyrophosphate. The presence of glucose causes a drop in the activity of adenylate cyclase and hence a drop in cyclic AMP levels. Glucose must be transported for this to happen, but it does not need to be broken down and metabolized. Non-metabolizable analogs of glucose, such as 2-deoxyglucose, cannot be degraded but can be transported and also cause catabolite repression.
Experimental Method: Beta-Galactosidase Assay
DAY 11 - Lab Period: Outline of Experiment
You will assay b-galactosidase from cultures grown under a variety of conditions to illustrate the regulation of the lac operon. Lactose and IPTG will be used as inducers and glucose will be added for catabolite repression.
Materials for cell growth
Klett colorimeter with green filter
Side arm flasks (sterile), 1 per student
Pipettes (sterile; 1, 5, 10 ml)
Vortex mixer
Tryptone Broth (contains, per liter, 10g tryptone & 5g NaCl)
Stock solutions of glucose (10%) and lactose (10%)
IPTG solution (25 mM) – keep refrigerated
Materials for b-galactosidase assay:
Disposable culture tubes (13 mm) for running b-galactosidase assay
Spectronic 20 for reading b-galactosidase assay
Bench top centrifuge such as GLC2
ONPG (4 mg/ml)
Sodium dodecyl sulfate (SDS) (0.1%)
Chloroform
Na2CO3 solution (1.0 M)
Medium E
Medium E (Vogel and Bonner, 1956) in g/L
K2HPO4, anhydrous.............. 10.0
NaNH4HPO4.4H2O.............. 3.5
Citric Acid.1H2O................... 2.0
MgSO4.7H2O........................ 0.2
You will be provided with a culture of a wild type E. coli strain (W1485) grown in tryptone broth medium. Dilute 2 ml of this culture into 22 ml fresh tryptone broth medium. Cultures should start out at approximately 20 Klett units. If too dilute, add more inoculum.
Set up 5 different cultures and make the following additions:
a) No additions
b) Lactose (final concentration 0.4%) add 1.0 ml of 10%
c) IPTG (final concentration 1.0 mM) add 1.0 ml of 25 mM
d) Glucose (final concentration 0.4%) add 1.0 ml of 10%
e) IPTG plus Glucose
Each student will grow one culture. The TAs will allocate cultures so that the class grows several of each type. Each pair of students will assay samples from one culture of each of the five types. Grow these cultures until they are between 75 and 100 Klett units (Cultures must double at least twice before assay). Record the Klett units and then carry out duplicate beta-galactosidase assays for each culture.
Principle of Beta-Galactosidase Assay
Beta-galactosidase is an enzyme which hydrolyzes beta-D-galactosides. It can easily be measured with chromogenic substrates (colorless substrates which when hydrolyzed yield colored products). An example is o-nitrophenyl-beta-D-galactoside (ONPG). This compound is colorless, but in the presence of beta-galactosidase it is converted to galactose and o-nitrophenol. The latter compound is yellow and can be measured by its absorption at 420 nm. If the o-nitrophenyl-beta-D-galactoside (ONPG) concentration is high enough, the amount of o-nitrophenol produced is proportional both to the amount of enzyme present and to the time the enzyme reacts with the ONPG. In order for the assay to be linear, the ONPG must be in excess. For best results, the amount of enzyme should be such that it takes between 5 minutes and 6 hours for a faint yellow color to develop – 20 minutes is convenient. The reaction is stopped by adding a concentrated Na2CO3 solution, which shifts the pH to 11. At this pH beta-galactosidase is inactive.
Assay of b-Galactosidase
Modified for Reading in the Spectronic 20. Instructions for using are on the machines.
a) Into each assay tube put:
1.0 ml of Medium E
0.2 ml of cells (use less cells and more medium E if the b-galactosidase activity is high; if necessary use a 10-fold dilution of your cells)
1 drop of 0.1% SDS (sodium dodecyl sulfate)
1 drop of chloroform
b) Vortex for 10 sec. to disrupt the cells.
c) To each tube add 0.2 ml of ONPG (4 mg/ml) solution.
d) Incubate tubes at 37° until sufficient yellow color has developed (A420 = 0.2-0.7). Estimate this by eye - a light but definite yellow color is needed. Keep track of time for each tube. Generally, 15 to 30 minutes is suitable.
e) Stop the reaction by adding 1 ml of 1 M Na2CO3 solution.
f) Spin the tubes in the low speed bench-top centrifuge for 5 min.
g) Read the Absorbance at 420 nm (= A420) in the Spectronic-20.
h) Use as a blank an assay containing 0.2 ml of medium E instead of a cell sample.
Calculations
a) Convert the absorbance at 420 to micromoles of o-nitrophenol by multiplying by 222.2 (this is based on a molar extinction coefficient of 4500 Mole-1 cm-1 for o-nitrophenol).
b) Divide by the time of incubation in minutes to get micromoles per minute.
c) Divide by volume of cells used, in ml, to get micromoles per minute per ml of cells.
(If cells were diluted, remember to multiply by the dilution factor.)
d) To compare the b-galactosidase activities of different bacterial cultures grown to different cell densities we calculate first the rate in terms of micromoles of o-nitrophenol produced, per minute, per ml of undiluted cells (steps a, b and c). Then multiply by 200/KU of culture. This gives the rate per 109 cells since a suspension of 109 cells/ml has a KU value of 200.
Rate (per 109 cells) = A420 x 222.2 x 200
Time (min) x Volume (ml) x KU
Experiment 5: SDS Polyacrylamide Gel Electrophoresis of Proteins
Perhaps the most widely used physical method in all of molecular biology is gel electrophoresis. The idea behind electrophoresis is that positive charges attract negative charges and vice versa. Conversely, two charges of the same sign repel each other. To perform electrophoresis we dissolve molecules which carry electrical charges in water and stick into the solution two electrodes - one positive and the other negative. When we switch on the current, the negatively charged molecules are attracted to the positive electrode and move through the solution until they reach it. The positively charged molecules move in the opposite direction. Molecules carrying charges are known as ions.
The greater the charge, the faster an ion will swim under the influence of an electrical attraction. On the other hand, the larger the molecule the more force needed to get it moving. Molecules of nucleic acid have exactly one negative charge for each nucleotide so these two factors cancel out. Consequently all molecules of DNA or RNA will move at the same speed towards the positive electrode as long as they are free in solution.
If we want to separate our DNA or RNA on the basis of size we must bring in an extra handicap to slow down the larger molecules - the gel. The gel is a meshwork of cross-linked polymer chains - usually of agarose for nucleic acids. The molecules of DNA are slowed down as they try to wriggle through the gaps in the gel meshwork. The larger they are, the harder it is to squeeze through the holes. The result is that a mixture of DNA molecules separates according to size, the smaller molecules moving through the gel much faster than the larger ones.
Nucleic acids all come with a built in negative charge, however proteins are not so convenient. Some of the amino acids from which proteins are built have a positive charge and some have a negative charge while most are neutral. So, depending on its overall amino acid composition, a protein may be positive, negative or neutral.
To avoid these complications we boil proteins in a solution of the detergent sodium dodecyl sulfate (SDS). Boiling destroys the folded 3D-structure of the protein, i.e. the protein is denatured. The SDS molecule has a hydrophobic tail with a negative charge at the end. The tail wraps around the backbone of the protein and the negative charge dangles in the water. The protein is unrolled and covered from head to toe with SDS molecules which give it an overall negative charge. What's more, the number of negative charges bound is proportional to the length of the protein.
To fully separate and unfold the individual polypeptide chains, we also need to break any disulfide bonds between cysteine residues. This is done by including an excess of the reagent b-mercaptoethanol when the proteins are boiled.
HO-CH2CH2-SH + HS-CH2CH2-OH + Protein-S-S-Protein
ß
HO-CH2CH2-S-S-CH2CH2-OH + Protein-SH + HS-Protein
The sample buffer also contains a tracking dye, in our case, bromophenol blue. This is smaller than the proteins and so moves ahead of them. When the bromophenol blue band reaches the end of the gel, we switch off.
So now we can separate proteins according to their sizes by running them through a gel. Because proteins are a lot smaller on average than DNA or RNA, we normally use a gel made of the artificial polymer, polyacrylamide, which gives smaller gaps in its meshwork than agarose.
This modified technique is known as sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and is often used to separate or purify proteins. It was first worked out by Laemmli (Laemmli, U.K., Nature, 227:680, 1970).
Polyacrylamide is made by polymerizing acrylamide, which is a potent neurotoxin. Avoid contact with unpolymerized acrylamide!! Bis-acrylamide is used to cross-link the polymer so making a meshwork. The polymerization of acrylamide plus bis-acrylamide is driven by the generation of free radicals. TEMED (N N N' N' - tetramethylethylene diamine) and ammonium persulfate are used to generate the free radicals. The base TEMED catalyses the decomposition of persulfate to give sulfate radical ions.
Once we have run our gel, we must stain it to see the protein bands. The two favorite choices are Coomassie Blue, a blue dye which binds strongly to proteins, or silver compounds. Silver atoms bind very tightly to proteins and yield black or purple complexes. Silver staining is more sensitive and more expensive.
To estimate the molecular weight of a protein of interest, we run a series of standard proteins of known molecular weight on the same gel. When the distance that each protein has run is plotted against the logarithm of its molecular weight we get a straight line. The distance run by the unknown protein is also plotted and the log of its molecular weight is read from the plot.
Experimental Method: Polyacrylamide Gel Electrophoresis
DAY 13 - Lab Period
You will grow cultures of different strains of E. coli and prepare samples for running on SDS-PAGE gels. You should be able to see some of the most plentiful proteins in E. coli. Each group will use one of the following sets of strains:
Set I - To illustrate the envelope proteins OmpF (38.3 Kd) and OmpC (37.1 Kd). These are two of the most plentiful proteins in the E. coli cell and give two of the most prominent bands in extracts of whole cells.
a) MC4000 parent c) TK821 ompR::Tn10 (tetracycline resistant)
b) SG480D76 D(malT-envZ) d) MH450 ompF::Tn5 (kanamycin resistant)
Set II - To illustrate the AdhE protein (96 Kd) - one of the largest proteins in E. coli. This can be seen near the top of the gel as there are very few proteins in this region. The adhE gene is induced anaerobically. Wild type cells will show little or no AdhE when grown aerobically. The adhC mutants express the adhE gene even in air.
a) DC271 wild type b) SHH31 DadhE
c) DC272 adhC d) DC430 adhC adhR
Preparation of Protein Samples for Loading onto Gel
a) Grow cultures of each strain in rich broth (the TA's will inoculate these the night before).
b) Adjust culture to 100 KU with sterile saline before using.
c) Take a 1.0 ml sample of cells from each culture and centrifuge it at 7000 x g for 2 min in an Eppendorf micro-centrifuge.
d) Discarded the supernatant and resuspend the cells in 400 ml of sample buffer.
e) Place the samples in a boiling water for 5 min
f) Load 15 ml of each sample onto a 10% SDS-PAGE gel along with a ladder made up of proteins of known MW. (If you have enough space on your gel, load two samples, of 10 and 20 ml, for each strain.)
Running the Gels
Before pouring the gel assemble the gel plates carefully to avoid leaks. Press down the smaller plate and clamp it. Push down when pouring the gel to ensure tight seal.
Mix the ingredients in the order given under “11% Running gel” (next page).
Note: the gel starts to set when you add TEMED and ammonium persulfate. The ammonium persulfate (AP) is a radical initiator and starts the polymerization.
[Before adding TEMED and AP, de-aerate the rest of the gel mix under low vacuum - this is not really necessary unless your gel mixture has been shaken up and looks bubbly.]
Pour the running gel carefully (bubbles disturb proper running).
[Cover the gel surface with water saturated n-butanol (avoid disturbing gel). Can omit this step if the gel will be used soon.]
Allow to set (about 30 minutes).
Remove water/n-butanol layer if present. Rinse top of gel carefully with distilled water.
Pour the stacking gel on top of the running gel. Put comb in position.
Allow to set (about 20 minutes).
Assemble gel apparatus.
Remove comb from gel. Load samples. Remove bubbles at bottom of gel by squirting buffer with a bent syringe.
Run at 15 milliamps per gel until the tracking dye reaches bottom - about 1.5 to 2 hours.
(Need 8 volt/cm for stacking gel and 15 volt/cm for running gel. For our apparatus 75-125 volts is about right.)
SWITCH OFF POWER!! Remove gel carefully, cut off a corner to indicate the orientation.
Stain overnight in Coomassie Blue.
DAY 14 - Unscheduled Lab:
Destain until bands are visible (two or three changes of destain solution).
Note: If your gel curls up during destaining, it may be uncurled by rehydration in 10% acetic acid (without methanol).
DAY 15 - Lab Period
Photograph or photocopy the gel and identify the important bands.
Measure the distance of migration of the protein bands of the protein standards of known molecular weight. Measure from top of running gel to leading edge of band.
Make a plot of migration distance versus log molecular weight for the standard proteins. (Use semi-log graph paper and plot MW on the log scale.) Use this to estimate the MW of important bands in experimental samples.
Gel Electrophoresis Solutions:
a) Sample Buffer
Glycerol 10.0 ml
10% SDS 23.0 ml
b-Mercaptoethanol 5.0 ml
500 mM Tris, pH 6.8 8.3 ml
0.1% Bromophenol blue 12.0 ml
distilled water 41.7 ml
b) Stacking Gel (need approx. 10-20 ml/gel)
distilled water 8.67 ml
4x upper gel buffer 1.67 ml
40% acrylamide/bis 1.32 ml
TEMED 40 ml
10% ammonium persulfate 70 ml (fresh solution)
c) 11% Running Gel (need approx. 30-40 ml/gel)
distilled water 18.75 ml
4x lower gel buffer 10.0 ml
40% acrylamide/bis 11.25 ml
TEMED 45 ml
10% ammonium persulfate 145 ml (fresh solution)
d) 4X Upper Gel Buffer
Tris (500 mM final) 6.06 gram
SDS (10%) 4 ml
in 100 ml of water
Adjust pH to 6.8 with HCI
e) 4X Lower Gel Buffer
Tris (1.5 M final) 18.1 gram
SDS (10%) 4 ml
in 100 ml of water
Adjust pH to 8.8 with HCI.
f) 40% Acrylamide/bis Solution
Get this from Fisher already made up - Note: Store in dark and cold.
g) Electrode Buffer
Tris 3.0 gram
Glycine 14.4 gram
SDS 1.0 gram
in 1000 ml
Do not adjust pH (it should be 8.3).
h) Stain
TCA 5.0%
Acetic Acid 10.0%
Methanol 10.0%
Coomassie Blue R250 0.1%
i) Destain
Methanol 30.0%
Acetic Acid 10.0%
Standard proteins
Standard proteins are Sigma Markers M-3788 (high MW range) and M-3913 (low MW range). Each vial was reconstituted with 100 µl of dH2O and contains approx 2-3.5 mg of protein per ml in Tris (62 mM, pH 8.0), 1 mM EDTA, 3% sucrose, 0.5% dithiothreitol, 2% SDS and 0.005% bromophenol blue. Use 5 ml per gel slot. [Note: marker mixtures may change from year to year – check with your TA.]
Standard proteins Mol Wt High Low
(M-3788) (M-3913)
Myosin, rabbit muscle 205,000 X
b-Galactosidase, E. coli 116,000 X
Phosphorylase b, rabbit muscle 97,000 X
Fructose-6-phosphate kinase, rabbit muscle 84,000 X
Bovine serum albumin 66,000 X X
Glutamic dehydrogenase, bovine liver 55,000 X
Ovalbumin, chicken egg 45,000 X X
Glyceraldehyde-3- dehydrogenase, rabbit muscle 36,000 X X
Carbonic anhydrase, bovine erythrocytes 29,000 X
Trypsinogen, bovine pancreas 24,000 X
Trypsin inhibitor, soybean 20,000 X
a-Lactalbumin, bovine milk 14,200 X
Aprotinin, bovine lung 6,500 X