be cell microbio script.7-37

Cell Biology & Microbiology Laboratory Course

FORMULAS AND HELPFUL TIPS

(I) CONCENTRATION CALCULATIONS

(1)

Example: 1 µL sample + 9 µL water (diluent) = 10 µL total volume → Dilution factor = 10 1 µL solution (dilution factor 10) + 1 µL water (diluent) = 2 µL total volume

→ Dilution Factor = 20 (2 x 10) 5 µL water (diluent) + x µL sample = 5 + x µL total volume

Reduction of concentration should be = 6 → 6 = 1 part of sample + 5 parts of dilution (all parts are equal in volume) → 5 equal parts of dilution = 5 µL water (diluent) / 5 = 1 µL → All 6 parts have the volume of 1 µL x µL sample = 1 µL

Conc. Reduction = initial volume of sample : total volume of the diluted solution = parts of sample : all parts of the dilution

(2)

Example: 1 µL sample + 9 µL water (diluent) = 10 µL total volume → Reduction of concentration = 1 : 10

1 µL 1:10 solution + 1 µL water (diluent) = 2 µL total volume → Reduction of concentration = 1:20 (1:2 x 1:10)

5 µL water (diluent) + x µL sample = 5 + x µL total volume Reduction of concentration should be = 1:6 → 1:6 = 1 part of sample + 5 parts of diluent (all parts are equal in volume) → 5 equal parts of dilution = 5 µL water (diluent) / 5 = 1 µL → All 6 parts have the volume of 1 µL x µL sample = 1 µL

(3)

C1 x V1 = C2 x V2 (4) V1 = Volume of solution in the

mixture

C1 = Concentration of the solution

C2 = Wanted concentration of the end-solution

V2 = Wanted Volume (volume

of all parts) Example: Concentration of the solution: C1 = 20 mg/mL Wanted concentration: C2 = 10 mg/mL Wanted volume: V2 = 5 mL C1 x V1 = C2 x V2 => 20 mg/mL x V1 = 10 mg/mL x 5 mL V1 = (10 mg/mL x 5 mL) / 20 mg/mL V1 = 2,5 mL 2,5 mL solution (20 mg/mL) + 2,5 mL dilution = 5 mL solution (10 mg/mL)


(II) PHOTOMETRIC MEASUREMENTS

DNA and RNA absorbs in the ultraviolet range, at a wavelength of about 260 nm because of the nitrogenous bases

Proteins absorb at 280 nm

Beer-Lambert Law: A = εcl A = absorbance value (no units) ε = extinction coefficient (constant for each substance)

=0.027 (μg/mL)-1 cm-1 for ssDNA =0.020 (μg/mL)-1 cm-1 for dsDNA =0.025 (μg/mL)-1 cm-1 for ssRNA

c = concentration of substance (units for DNA/RNA = μg/mL) l = light path length

= 1 cm for standard cuvettes For the light path length in 96 well plates use the formula

4 ∗ ∗

V = sample volume d = mean diameter of the well Does not account for the meniscus of the liquid

= 0.29 cm for a 96 well plate with a 100 μL sample volume = 0.56 cm for a 96 well plate with a 200 μL sample volume

(5)

Example 10 µL ssDNA + 90 µL water mixed in the cuvette (Conc. Reduction = 1:10) Spectrophotometric measured absorbance (OD) of ssDNA A = 0.9 Measured blank (10 µL buffer of ssDNA + 990 µL water) A = 0,225

ssDNA absorbance without the background of solutions: A ssDNA – A blank ssDNA A = 0.9 – blank A = 0,225 → A = 0.675

0.675 = 0.027 ⁄ ∗

x c x 1 cm x (Concentration Reduction)

c = . ∗

. ∗μ ⁄

c = 250 µg/mL The original 10 µL ssDNA has a concentration of 250 µg/mL

The ratio of absorbance at 260 to 280 nm (A260/A280) should be In case of DNA: 1.8 (>1.75) In case of RNA: 1.8 - 2.1

(6)


(III) UNITS

Unit prefix - indicate multiples or fractions of the units

tera giga mega kilo Unit

T G M k

= 1012

= 109 = 106 = 103

= 0

Unit Milli Micro Nano Pico

m µ n p

= 0 = 10-3

= 10-6 = 10-9 = 10-12

103 = 1,000 106 = 1,000,000 109 = 1,000,000,000 1012 = 1,000,000,000,000

10-3 = 0.001 10-6 = 0.000001 10-9 = 0.000000001 10-12 = 0.000000000001

Solids in liquid 1 g/L = 1 mg/mL = 1 µg/µL = 1 ng/nL = 1 pg/pL 1 g/L = 0.001 g/mL = 0.001 mg/L 1 mg/mL = 1000 g/mL = 1000 mg/L

(IV) GENERATION TIME

Generation time is per definition the time interval required for the cells (or population) to divide:

G tn

G = generation time t = time (in minutes or hours) n = number of generations (number of times the cell population doubles during the time interval)

(7)

Number of bacteria B = number of bacteria at the beginning of a time interval b = number of bacteria at the end of the time interval

b Bx2 ↔

n = 3.3 log(b/B)

This equation is an expression of growth by binary fission

(8)

G t

3.3log bB

(7)+

(8)


(V) CELL NUMBER WITH HEMOCYTOMETER

1 mm3 = 1 µL (9)

Thoma Chamber Size of the smallest square: 0.0025 mm² x 0.1 mm Depth = 0,00025 mm³ = 0,00025 µL Size of the next bigger square: Next bigger square = 16 x smallest square 16 x 0.0025 mm² x 0.1 mm Depth = 0.004 mm³ = 0.004 µL

1. Count the cells in a square of your choice 2. Calculate the cell number to the standard unit cells per mL 3. If the sample was diluted, multiply the calculated cell concentration by the

dilution factor

Example 12 cells are in the next bigger square (0.004 µL) 12 cells per 0.004 µL = 12 cells/0.004 µL Use the rule of three

3000 000 cells/mL The sample was diluted (1 part + 4 parts dilution) with a dilution factor of 5 3000 000 cells/mL x 5 = 15000 000 cells/mL

The smaller the volume of interest- the smaller the number of cells Example of the same concentration 2 points in the smaller dotted square (1/4 the size of the big one) 8 points in the big square (4 times more than in the small one)

Example of a dilution 2 points in the smaller dotted square (1 part) Transfer into a big square filled with dilution solution (3 parts) Concentration Reduction (2) = 1:4

Calculate the concentration of the dilution Original concentration: 8 points in the big square, dilution 1:4 8 x ¼ = 8 4 = 2 points in the dilution


(VI) TRANSFORMATION EFFICIENCY

Transformation efficiency # of colonies x 106pg x volume of transformants pg plasmid DNA 1 µg X µL plated

# of colonies = colonies counted in the petri dish pg plasmid DNA = used amount of pUC19 DNA for the tansformation in pg 106 pg / 1 µg = Calculation from the unit pg to the unit µg volume of transformants = total amount of the mixture (in µL) before X µL were taken out to plate it X µL plated = amount of liquid, which was given on agar in the petri dish (in µL)

(10)

Example 30 successfully transformed blue colonies are on the agar 300 pg Bluescript plasmid DNA were used 1053 µL were mixed as a volume of transformants

50 µL competent cells 3 µL Bluescript DNA

1000 µL LB medium 200 µL were spread

30 colonies x 1000000 pg x 1053 µL transformants 300 pg Bluescript DNA 1 µg 200 µL plated

30 colonies x 106 pg x 1053 µL 526500 colonies 300 pg Bluescript DNA 1 µg 200 µL µg Bluescript DNA

In this example, 526,500 colonies were transformed per µg Bluescript plasmid.

=

Unit 1 Cell Biology & Microbiology Laboratory Course

UNIT 1. ACCURATE PIPETTING OF LIQUIDS, SERIAL DILUTIONS AND STERILE TECHNIQUE This unit aims to introduce you to a number of crucial skills to be able to succeed in the laboratory. These skills include accurate pipetting of liquids, preparation of serial dilutions, data analysis and sterile technique.

I. ACCURATE PIPETTING OF LIQUIDS (PART I) Answer these following questions before you join the practical course. 1.1 Name the three main types of pipettes.

1.

2.

3.

1.2 Name the volume ranges of the three micropipettes: Micropipette Range in µL

P20

P200

P1000

1.3 In which situations do you have to use the forward technique and in which the reverse technique?

Technique Sort of liquid

Forward technique

Reverse technique


INTRODUCTION The first part of today´s session is dedicated to give you experience in using various types of pipettes usually found in a research lab. There are three main types of pipettes used in the lab to move liquids from one reservoir to another: (i) Pasteur pipettes, (ii) volumetric or serological pipettes, and (iii) micropipettes.

Figure 1. Three main types of tools used in the laboratory to move liquids. (i) Pasteur pipettes, (ii) pipettes, and (iii) micropipettes.

(i) Pasteur pipette: small, tapered glass tube, not graduated, and used with a bulb. It is used to dispense liquid when the volume is not critical. (ii) Volumetric or serological pipette: glass or plastic, calibrated to deliver any amount in the graduated scale from 1 to 25 mL, and used with a bulb or Pipette-Aid. A Pipette-Aid is an electric pipettor that has two buttons which control the flow of liquid in and out of the pipette. Pipettes are available that hold 1 mL, or 2 mL, or 5 mL, or 10 mL, or 20 mL. The proper pipette must be chosen for the task at hand. For example, a 1 mL pipette will more accurately measure 1 mL than will a 5 mL pipette. Thus, you should choose the smallest volume pipette that will do the job. (iii) Micropipette: used with plastic pipette tips, calibrated to dispense smaller volumes ranging from 0.5 µL to 1000 µL. There are different micropipettes from different brands designed for a particular range of volume. The volume capacity is indicated at the top near the plunger. The micropipettes available in the lab cover three volume ranges: 2- 20 µL (p20), 20- 200 µL (p200), and 100- 1000 µL (p1000).


You must choose the correct pipette for the volume to be measured. The lowest end of each volume range is subject to the greatest percentage error. For accuracy, you must stay within the calibrated range of each micropipette: 2- 20 µL (p20), 20- 200 µL (p200), and 100- 1000 µL (p1000). Do not push the micropipette outside of the indicated volume range. If you do so, this will lead to inaccurate measurements and damage to the pipette. For each of these micropipettes, a different tip is used. (iii a) Micropipette overview

Figure 2. Moving the button/piston displaces air which moves the liquid. 1. The piston moves to the appropriate position when the volume is set. 2. When the operating button is pressed to the first stop, the piston expels the same volume of air as indicated on the volume setting. 3. After immersing the tip into the liquid, the operating button is released. This creates a partial vacuum and the specified volume of liquid is aspirated into the tip. 4. When the operating button is pressed to the first stop again, the air dispenses the liquid. To empty the tip completely the operating button is pressed to the second stop (blow out).

(iii b) Pipetting Technique: Forward versus Reverse The forward technique is the standard for pipetting aqueous liquids, whereas the reverse technique is used for pipetting viscous solutions or liquids with a tendency to foam.


Figure 3. Forward versus reverse pipetting technique. Forward pipetting modus: 1. Press the operating button to the first stop. 2. Dip the tip into the solution to a depth not more than 3-4 mm, and slowly release the operating button. Wait 1-2 seconds and withdraw the tip from the liquid, touching it against the edge of the reservoir to remove excess liquid. 3. Dispense the liquid into the receiving vessel by gently pressing the operating button to the first stop and then press the operating button to the second stop. This action will empty the tip. Remove the tip from the vessel, sliding it up the wall of the vessel. 4. Release the operating button to the ready position. Reverse pipetting modus: 1. Press the operating button to the second stop. 2. Dip the tip into the solution to a depth not more than 3-4 mm, and slowly release the operating button. This action will fill the tip with a volume that is larger than the set volume. Wait 1-2 seconds and withdraw the tip from the liquid. 3. Dispense the liquid into the receiving vessel by pressing the operating button gently and steadily to the first stop. This volume is equal to the set volume. Hold the button in this position. Some liquid will remain in the tip, and this should not be dispensed. 4. The liquid remaining in the tip can be pipetted back into the original solution or disposed of together with the tip. 5. Release the operating button to the ready position.

(iii c) Preventing cross-contamination Pipette-to-sample: A contaminated pipette, or its tips, can contaminate your samples. You can avoid such contamination by wiping your pipette with 70 % ethanol, by using sterilized tips, tips with filters, and by changing the tip after pipetting each sample. Sample-to-pipette: Samples or aerosols can enter the cone of the pipette. You can avoid such contamination by keeping the pipette vertical when pipetting in order to prevent liquid from running into the pipette body, by releasing the push-button slowly, by using filter tips, and by storing the pipette vertically. Sample-to-sample: The remains of sample A can mix with sample B inside the tip and may cause a false result. You can avoid such contamination by changing the tip after each sample. (iii d) Calibration of micropipettes (accuracy versus precision) Calibration of micropipettes means determining the difference between the dispensed volume and the selected volume (accuracy). Precision refers to the repeatability of the pipetting. It is expressed as standard deviation (SD) or coefficient of variation (CV). What is needed is both, accuracy and precision.


Figure 4. Accuracy versus precision. When the set volume is for example 20 µL, accurate but not precise: The mean volume is the correct (set) volume but individual pipettings differ from the set volume. Precise but not accurate: There is not variation between the separate pipettings, but the mean volume differs from the set volume. Accurate and precise: The mean volume is the set volume and there is no variation between different pipettings.

(iii e) Factors affecting micropipette’s accuracy Factors affecting micropipette’s accuracy include: Position: Pipettes should be held vertical during the aspiration of liquids; however, some users often hold pipettes at many different angles during a pipetting interval. Holding a pipette 30 º off vertical can cause as much as 0.7 % more liquid to be aspirated due to the impact of hydrostatic pressure. Tips: To ensure accurate results, use tips specified by manufacturers. If using other tips, the properties should be tested (proper hold, dispensing of the correct and whole volume). Release of Plunger: Releasing the plunger abruptly can cause liquid to accumulate inside the instrument which in turn can be transferred to other samples causing variability in sample volume and is a potential source for cross-contamination. It is recommended that a smooth, consistent pipetting rhythm is employed since it helps to increase both accuracy and precision. Immersion Depth: The pipette tip should only be inserted into the vessel containing the liquid to be transferred about 3-4 mm. Environmental conditions: Temperature, air pressure, and humidity are the main error sources from the environment. The greatest contributor to error is liquid temperature (see Figure 5). Density: Density is the mass/volume ratio of the liquid. The density varies with the temperature and air pressure. The density of water is at room temperature (20-25 °C) 0.996 g/mL.


Figure 5. Impact of liquid temperature and density on pipetting accuracy.

MATERIALS Deionized distilled H2O (ddH2O in 100 mL Flask) Tween-20 (in 100 mL Flask) Pasteur Pipette 5, 10 and 25 mL Pipette Pipette-Aid Set of micropipettes (P20, P200 and P1000) Set of tips 100 mL beaker Reaction tubes (1.5 mL microcentrifuge tube and 15 mL reaction tube) Rack for microcentrifuge tubes and reaction tubes Analytical balance EXPERIMENTAL PROCEDURE (i) Macropipetting with a Pasteur pipette: When you need to take a liquid, but do not need a specific volume, you can use a Pasteur pipette. 1.4 Using this pipette, fill in the 15 mL reaction tube with H2O from the 100 mL flask.

How many times did you need to pipette in order to fill in the 15 mL reaction tube?

(ii) Macropipetting with a pipette and a Pipette-Aid: You will practice dispensing different volumes of liquid using the 5, 10 and 25 mL plastic disposable pipette. Using this pipette, practice pipetting with the Pipette-Aid. Place the tip of the

pipette just deep enough into solution to obtain the volume of solution needed but do not hit the bottom of the beaker (or any reservoir). Get used to the rate at


which the fluid is drawn up and expelled. The rate varies depending on how hard you push the buttons. Expel the ddH2O into an empty beaker.

Pay attention on the pipette calibration markings. For example, as the 10 mL pipette is filled from the tip, the volume it contains when the bottom of the meniscus is at the “7 mL” mark is actually 3 mL. Thus, the markings indicate the volume of solution that has been let out of the pipette.

Learn how to insert the pipet into a flask without touching the sides and hold it there while withdrawing the liquid.

1.5 Draw up the following volumes: 1, 2.5, 3, 5, 10.5, and 20 mL. Which pipette did you use to achieve the most accurate measurement of the volume needed? Explain why.

(iii) Micropipetting using a micropipette: A micropipette is used to dispense small volumes (< 1 mL) of liquid. In this experiment you will practice working with three micropipettes of different volume ranges. Take the P20, P200 and P1000 micropipettes and look at the “digital volume

indicator”. The P20 has 3 display spaces: The top space reads tens of µL The middle space reads ones of µL The bottom space reads tenths (in red) of µL The P200 has 3 display spaces: The top space reads hundreds of µL The middle space reads tens of µL The bottom space reads ones The P1000 has 3 display spaces: The top space reads of thousands of µL

The middle space reads hundreds of µL The bottom space reads tens of µL Once you have understood the volume display, take the P1000 micropipette to

measure 500 µL ddH2O and deliver it into a microcentrifuge tip. Set the volume and make sure that the volume you choose is within the range of

the micropipette. Load the correct tip onto the micropipette by firmly pushing the barrel into the tip

which is still in the pipette-tips box. Tap it up and down to make sure it is seated. Before you draw up the liquid from its reservoir, check that you are using the right

micropipette and doublecheck if the set volume is correct. Draw up the liquid by using the “forward pipetting technique” mentioned above. Expel the ddH2O into an empty microcentrifuge tube and deposit the tip directly

into the sharps container by pressing the ejector button. Practice with P20 and P200 pipettes until you get used to them.

1.6 Draw up the following volumes of ddH2O: 12.5 µL, 25 µL, 200 µL, 650 µL,

666 µL, 970 µL, and 1300 µL. Expel the ddH2O into an empty microcentrifuge


tube. Which pipette did you use to achieve the most accurate measurement of the volume needed? Explain why.

1.7 Take the P200 and the 1.5 mL reaction tube, which contains 30 µL water and pipette 20 µL out of it. Use the P200 again together with the 1.5 mL reaction tube, which contains 190 µL water and pipette 180 µL out of it. Is there any difference between the positions of the first stops?

1.8 Try to find out the amount of water inside of the 1.5 mL reaction tube labeled with X-number. Look at the liquid and the scale on the side of the tube, to get an idea of the best fitting pipette. Take up the liquid into the tip. If there is any liquid left in the tube, dispense the water carefully and increase the numbers on the digital volume indicator. Try again to take up the liquid. If there are bubbles inside of the tip, decrease the numbers on the digital volume indicator (Attention! Please check the pipette’s range before using it!). Take another pipette with a different range, if necessary. Get closer to the actual amount of water with every attempt and note your result.

1.9 You will now practice the “reverse pipetting technique” by drawing up 500 µL Tween-20. What did you observe?

1.10 Name the different steps for pipetting with the micropipette. Mind the correct order:

Step Legend 1.

2.

3.

4.

5.

6.

7.

8.


9.

10.

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1.11 Name the don’ts during pipetting with the micropipette: DON‘TS Legend


II. ACCURATE PIPETTING OF LIQUIDS (PART II) Answer the following questions before you join the practical course. 1.12 Mention the density of water. 1.13 What does BSA means and what is it? 1.14 Explain the principle of a standard curve. 1.15 Calculate the concentration reduction (2) and the dilution concentration of each

tube in the serial dilution in Figure 7. initial sample concentration

initial sample volume

Diluent volume

Total volume

Concentration reduction (2)

Dilution concentration

2 mg/mL 500 µL 500 µL 500+500= 1000 µL

500 : 1000 = 1 : 2 = ½

½ x 2 mg/mL= 1 mg/mL

1 mg/mL

(iv) Micropipette calibration: In this exercise you will calibrate your P1000 micropipettes which will tell you how accurate your measurements are. You will pipette ddH2O into a number of microcentrifuge tubes. The amount of water dispensed, and therefore the accuracy with which you pipette, will be determined by weighing. Take five sets of three microcentrifuge tubes, label them (A1-3 to E1-3) and weigh

each tube Using a P1000 pipette, dispense volumes of 0.2 mL to each tube in set “A”,

0.4 mL to each tube in set “B”, 0.6 mL to each tube in set “C”, 0.8 mL to each tube in set “D”, and 1 mL to each tube in set “E”

1.16 In your computer use a spreadsheet to make the following calculations: - Substract the two weights for each test to determine the volume dispensed into

each (assume that the density of H2O is 1.0 g/mL) - Determine the mean and standard deviation for each set of triplicates done - Plot the “volume dispensed” (Y) versus the “micropipette setting” (X) on a line

graph - Calculate the best fit for this data using linear regression and add the fit to the

line graph. See the following example:


Figure 6. Example of visualized P1000 Micropipette calibration results. Volume dispensed (Y) is plotted versus the micropipette setting (X).

DISCUSSION 1.17 What volume is being measured?

The charts below show the type of micropipette being used and its “digital volume indicator” setting. What volume is being measured in each case?

Type Settings Volume measured (µL) P1000

0 1 0 0 0 0 1. 4.

2 0 0 7 5 9 2. 5.

5 0 7 0 3 2 3. 6.

Type Settings

Volume measured (µL)

P200

1 0 0 1 1 0 1. 4.

2 1 2 0 9 9 2. 5.

0 2 1 2 9 9 3. 6.

Type Settings

Volume measured (µL)

P20

0 2 1 0 0 1 1. 4.

2 0 9 0 5 3 2. 5.

0 0 9 1 7 4 3. 6.


1.18 Calculate accuracy and precision of your P1000 micropipette when measuring 200 µL ddH2O according to the following formulas:

III. SERIAL DILUTIONS INTRODUCTION A common procedure in molecular cell biology, microbiology and biochemistry is the quantitative determination of some compound in a sample. In most cases a colorimetric assay is used for such determinations. Colorimetric assays involve the use of a spectrophotometer to determine the absorbance of light due to the presence of a colored analyte. The quantitative aspects of these assays are based on Beer-Lambert´s Law which states that the concentration of a colored compound in a solution is directly proportional to the absorption of light by the solution. A = ε l c A = absorbance ε = molar absorptivity (varies with the wavelength of light used in the measurement) I = path length c= analyte concentration In this part you will use an assay (“Bio-Rad protein assay”, Bio-Rad Laboratories, Munich, Germany) to measure protein concentration. The Bio-Rad Protein Assay is a dye-binding assay in which a differential color change of a dye occurs in response to various concentrations of protein (1). The absorbance maximum for an acidic solution of Coomassie® Brilliant Blue G-250 dye shifts from 465 nm to 595 nm when binding to protein. The Coomassie blue dye binds primarily to basic and aromatic amino acid residues (2). Today you will prepare solutions containing various concentrations of a protein (Bovine Serum Albumin, BSA) and read the absorbance of these solutions at


595 nm with a spectrophotometer. This will allow you to prepare a standard curve. Comparison of the absorbance of a sample to a standard curve will provide you a relative measurement of a sample´s protein concentration. LITERATURE 1. Bradford, M., Anal. Biochem., 72, 248 (1976). 2. Compton, S. J. and Jones, C. G., Anal. Biochem., 151, 369 (1985). MATERIALS BSA (2 mg/mL) Phosphate Buffer Saline PBS Coomassie® Brilliant Blue G-250 dye reagent (contains dye, phosphoric acid and

methanol) Sample (cell lysate) Set of micropipettes (P20, P200 and P1000) Set of tips Reaction tubes (1.5 mL microcentrifuge tube) Microtiter plates Spectrophotometer

EXPERIMENTAL PROCEDURE Using serial dilutions (1/2 serial dilutions) prepare BSA solutions in buffer (PBS)

with the following concentrations: 2, 1, 0.5, 0.25, 0.125 mg/mL. Prepare each of these solutions. Find the pipetting scheme below (Figure 7).

Figure 7. Pipetting scheme of a serial dilution with a dilution factor of 2, which leads to a 1:2 concentration reduction of the starting solution. One part of the starting solution (very right tube, concentration of 2 mg/mL) is diluted with one part diluent. It is mixed 1 Part : 1 Part. The concentration of the tube in the black box is to be determined with the help of the standard curve. Changed after: http://www.clker.com/cliparts/o/Z/8/K/a/X/eppi-lysat-b-hi.png,

1.19 Draw a pipetting scheme before you start pipetting.


Add 97 μL of diluted dye reagent to each well (A1 to A12, total = 12 wells) of a microtiter plate.

Pipet 3 µL of BSA solution 0.125 mg/mL to well A1 and A2; 3 µL BSA solution 0.25 mg/mL to well A3 and A4; 3 µL BSA solution 0.5 mg/mL to well A5 and A6; 3 µL BSA solution 1 mg/mL to well A7 and A8; 3 µL BSA solution 2 mg/mL to well A9; 3 µL sample solution of unknown concentration to well A10 and A11 and 3 µL PBS -as a blank- to well A12. Remember to change the pipette tips while using a new BSA concentration (to

avoid cross-contamination). Mix sample and reagent by repeated pipetting. Be careful not to produce air

bubbles while mixing. Incubate at room temperature for at least 5 minutes and measure absorbance at

595 nm. DISCUSSION 1.20 Why do we need to measure the absorbance of PBS? What is meant with PBS

as a blank? 1.21 Plot absorbance versus concentration to get a standard curve. 1.22 Use the standard curve to determine concentration of protein in the sample (cell

lysate). 1.23 Calculate the dilution factor according to formula (1) and complete the chart

below.

Dilutionfactortotalvolumeofthedilution

initialvolumeofstartersolutionallpartsofthedilutionpartsofstartersolution

(1)

Dilution Factor 50 mL starting solution in a final volume of 500 mL 10

200 mL starting solution in a final volume of 500 mL

250 mL in 500 mL

320 µL in 80 mL

2 Parts starting solution in a total of 5 Parts solution

50 mL starting solution mixed with 500 mL dilution solution 11

200 mL starting solution mixed with 500 mL dilution solution

250 mL with 500 mL

320 µL with 80 mL

2 Parts starting solution with 5 Parts dilution solution


1.24 Calculate the concentration reduction of the dilution according to formula (2) and complete the chart below.

Conc.Reduction initialvolumeofstartersolution:totalvolumeofthedilution partsofstartersolution:allpartsofthedilution

(2)

Concentration Reduction 50 mL starting solution in a final volume of 500 mL 50:500 = 1:10

200 mL starting solution in a final volume of 500 mL

250 mL in 500 mL

320 µL in 80 mL

2 Parts starting solution in a total of 5 Parts solution


50:550 = 1:11


250 mL with 500 mL

320 µL with 80 mL

2 Parts starting solution with 5 Parts dilution solution

1.25 We use a BSA stock solution with a concentration of 2 mg/mL, which means, that there are 2 mg BSA in 1 mL solution. The rule of three can be used to calculate the amount of BSA in 1 µL:

(3)

Calculate the amount of BSA or the amount of liquid BSA stock 2 mg / 1 mL BSA stock 2 mg / 1 mL

1 µg 0.5 µL 85 mg 42.5 mL

2 µL 10 mg

20 µg 270 µL

1.5 mL 7 µg

3.5 mg 1 mL

500 µL 14.4 µg

1 g 13 µL


IV. STERILE TECHNIQUE INTRODUCTION By using sterile techniques you can minimize the exposure of your solutions and cell cultures to contaminants (present in the air, hand and mouth while talking, etc.) that would interfere with your results and would even kill your cell cultures. Thus, it is very important to know how to work properly to keep your cells and solutions free of contaminants. Today, and in next lab course units, you will practice how to work in a cell culture hood. (i) Basic rules The working area should be as far away from traffic as possible. Set the working area up to minimize the time that sterile cultures and media are

open. That means: Clear the working area beforehand and have everything you will need ready at hand.

If you are using a cell culture hood, wipe it down with an antiseptic/cleaning agent before use.

You can only touch sterile items with other sterile items Hold caps from sterile bottles face side up in your hand when you uncap them.

Do not touch the sterile side of the cap. You are the most likely source of bacteria and fungi that could contaminate your

cultures. Talking close to an open culture is the surest way to contaminate it. Minimize the amount of open bottles and the time bottles are open and don’t talk while working with open bottles.

(ii) Common mistakes Pipetting up too far in the pipette. Touching the tip of the pipette against the outside of a bottle, the ground, bench

(anywhere not sterile). Dropping an opened container or tube to the ground. Talking into an open dish or bottle. (iii) Rules when working with a cell culture hood Although the danger of contamination is diminished in a cell culture hood, sterile technique is still essential in keeping your cultures contaminant free. In a cell culture hood, the major cause of contamination is movement of the arms in and out of the cabinet, which breaks the air curtain and disturbs the air flow required to prevent contamination. Check that the cell culture hood is on and the air is circulating. Do not block airflow by placing items on it. Do not bring more items into the cell culture hood than necessary. Wipe down the surface with 70 % EtOH before and after each use. Minimize hand movement in and out of the cell culture hood.


MATERIALS 70 % EtOH Cell culture Media Paper towels Wrapped 10 mL pipettes Pipette-Aid T25 Cell culture flask Gloves EXPERIMENTAL PROCEDURE Today you will learn how to dispense liquids from a bottle with cell culture media into a cell culture flask. After preparing your culture, you will place your cell culture flask in an incubator at 37 °C and let it incubate for a week. In next lab course unit, you will be able to check whether you worked properly by examining your cell culture for contaminants. Check that the cell culture hood is on and the air is circulating. Bring needed items into the cell culture hood. Wear gloves and wipe them with 70 % EtOH. Wipe down the surface with 70 % EtOH before use. Wipe down the cell culture media bottle surface with 70 % EtOH before use. Take a 10 mL pipette. Pull the plastic sleeve down and away from the pipette to

snap it open. Only pull it about 1/5 of its length. Insert the pipette into the Pipette-Aid. Hold the Pipette-Aid in one hand while you remove the pipette from the wrapping with your other hand (be careful not to touch the pipette to any surface as you do this).

Open the cell culture media bottle and place the cap face side up. Do not touch the sterile side of the cap.

With the pipette, draw up 8 mL of cell culture media without touching the outside of the bottle.

Open the cell culture flask and place the cap face side up. Do not touch the sterile side of the cap.

Expel the liquid into the cell culture flask without touching the outside of the bottle. Close both the cell culture flask and cell culture media without touching the sterile

side of the cap. Place the cell culture flask into the incubator. Remove items from the cell culture hood. Wipe down the surface with 70 % EtOH. Switch off the cell culture hood. LITERATURE 1. Barker, Kathy. At the bench: A Laboratory Navigator. Cold Spring Harbor

Laboratory Press, 2005.


UNIT 2. BASIC MICROBIOLOGY LABORATORY TECHNIQUES (PART I) The aim of the next two units is to introduce you to a number of basic techniques used to study microorganisms. These techniques include Gram´s staining to identify bacteria, bacterial culture methods, measurement of bacterial growth and growth limitation. In this unit you will learn to distinguish between gram positive and gram negative bacteria by following the Gram staining method. Gram staining is a valuable diagnostic tool in clinical settings as both bacterial types react to antimicrobial agents differently. Antimicrobial activity is measured by determining the smallest amount of agent needed to inhibit the growth of a test organism, a value called the minimum inhibitory concentration (MIC). In this section you will also learn how to determine the MIC-value of an antimicrobial agent. The correct handling of a light microscope In this unit you will work with a microscope. Microscopes are expensive scientific instruments. The Rhine-Waal University of Applied Sciences provides Zeiss microscopes. Handle them properly and carefully and they will last for many years! When moving your microscope, always carry it with both hands. Grasp the arm with one hand and place the other hand under the base for support.

Figure 8. Parts of the Microscope.

Turn the revolving nosepiece so that the lowest power objective lens (4 x magnification, labeled with the red ring) is "clicked" into position. Attention: Use the 100 x objective lens (white ring) only after consulting the lab instructors. There is the risk of a damage in case of incorrect use. Place the microscope slide on the stage and fasten it with the stage clips. You can push down the back end of the stage clip to open it. Look at the objective lens and the stage from the side and turn the focus knob so that the stage goes upward. Move it as far as it will go without touching the slide! Use the fine adjustment knob only, when the distance of the stage to the


objective goes under 0.5 cm. Now, look through the eyepiece and adjust the dimmer switch and diaphragm for the greatest amount of light. Slowly turn the coarse adjustment so that the stage goes down (away from the slide). Continue until the image comes into focus. Use the fine adjustment, if necessary, for fine focusing. Do not prepare your sample on the stage! The work with liquids on the stage is forbidden. Move the stage with the microscope slide around so that the image is in the center of the field of view and readjust the dimmer switch or diaphragm for the clearest image. When you are able to identify the image of your sample, change to the next objective lenses with only minimal use of the focusing adjustment. Use the fine adjustment. If you cannot focus on your specimen, repeat the previous steps with the higher power objective lens in place. Do not allow the objective lens to touch the slide! Do not touch the glass part of the lenses with your fingers. Use only special lens paper to clean the lenses. When finished, lower the stage, click the low power lens into position and remove the slide. Turn out the light before you unplug the microscope. Always keep your microscope covered when not in use. I. GRAM’S STAINING Answer these following questions before you join the practical course. 2.1 Name the bacteria which are examined.

1.

2.

2.2 Which microscopes objective lens is only to be used after consulting the lab instructors?

2.3 Sequence the steps of the experiment. If things happen at the same time give the same number. Number Steps of the experiment

Iodine is added

Decolorizer is added

1 Crystal violet solution is added to the bacteria

Insoluble crystal complex are formed

Safranin is added

2 Crystal violet solution enters the cells

Cells with peptidoglycan-poor cell walls get colorless

All bacteria are stained

Gram negative cells are stained

Gram positive cells are stained

Lipid layer from the gram negative cells are dissolved


INTRODUCTION The Gram staining method is one of the most important staining techniques in microbiology. It is almost always the first test performed for the identification of bacteria. In order to understand how staining works, you should know about the physical and chemical nature of stains. Stains are generally salts (a compound composed of a positively charged ion and a negatively charged ion) in which one of the ions is colored. For example, the dye methylene blue is actually the salt methylene blue chloride which will dissociate in water into a positively charged methylene blue ion which is blue in color and a negatively charged chloride ion which is colorless. Dyes or stains are divided into two groups: basic and acidic. If the color portion of the dye resides in the positive ion, as in the above case, it is called a basic dye (examples: methylene blue, crystal violet, safranin). If the color portion is in the negatively charged ion, it is called an acidic dye (examples: nigrosin, congo red). When using a basic dye, the positively charged color portion of the stain combines with the negatively charged bacterial cytoplasm and the organism becomes directly stained. An acidic dye reacts differently. Since the color portion of the dye is on the negative ion, it does not combine with the negatively charged bacterial cytoplasm. Instead, it forms a deposit around the organism, leaving the organism itself colorless. This type of staining is called indirect or negative as the organism is seen indirectly, and is used to get an accurate view of bacterial size and shapes. Today, we will make a direct stain of two bacterial strains by using the basic dye crystal violet as a primary stain. The bacteria that retain the crystal violet-iodine complex appear bluish purple under microscopic examination and are classified as gram positive bacteria. Others that are not stained by crystal violet are referred to as gram negative bacteria and appear pinkish red. Principle and interpretation Gram staining is based on the ability of bacterial cell wall to retain the crystal violet dye during solvent treatment. Gram positive bacteria have very thick cell walls consisting primarily of peptidoglycan whereas most of the cell wall in gram negative bacteria is composed of an outer membrane containing lipids, proteins and polysaccharides. Only 10 % of the gram negative total cell wall is made of peptidoglycans (Figure 9). When bacteria are incubated with crystal violet, dye enters into the cell. Iodine is subsequently added as a mordant to form an insoluble crystal violet-iodine complex. This step is commonly referred to as fixing the dye. Subsequent treatment with a decolorizer (a mixed solvent of ethanol and acetone) dissolves the lipid layer from the gram negative cells which enhances the leaching of crystal violet from the cells to the surroundings. In contrast, in gram positive cells, the solvent dehydrates the thicker peptidoglycan cell wall thereby closing the pores as the cell wall shrinks during dehydration. Thus, the diffusion of the violet-iodine


complex is blocked and bacteria remain stained. Finally, a counterstain of safranin is applied to the smear to give colorless-gram negative bacteria a pink color.

Figure 9. Schematic diagrams of gram-positive (a) and gram-negative (b) cell walls. The Gram stain photo in the center shows cells of Staphylococcus aureus (purple, gram-positive) and Escherichia coli (pink, gram-negative). Source: Figure 3.13 Brock Biology of Microorganisms 13/e.

MATERIALS Deionized distilled H2O (ddH2O in 100 mL Flask) Microscope slides Gram´s crystal violet solution Gram´s iodine solution Gram´s decolorizer solution Gram´s safranin solution Micropipette (P1000) and tips Bunsen burner Gram positive bacteria: Bacillus subtilis Gram negative bacteria: Escherichia coli

EXPERIMENTAL PROCEDURE (i) Prepare a slide smear Transfer a drop of the Bacillus subtilis suspended culture on a slide. It should only

be a very small amount of culture. Spread the culture with a pipette tip to an even thin film over a circle of 1.5 cm in

diameter. Since it is possible to put two small smears on a slide, transfer and spread a drop of Escherichia coli suspended culture into the same slide.

Allow to air-dry the slide and fix it over a gentle flame, while moving the slide in a circular fashion (three times, during 1 s/time) to avoid localized overheating. The applied heat helps the cell adhesion on the glass slide to make possible the subsequent rinsing of the smear with water without a significant loss of the culture.


(ii) Gram staining Flood the fixed smear with Gram´s crystal violet solution. Let stand for

60 seconds. Pour off the stain and gently wash with water. Flood with Gram´s iodine solution. Allow it to remain for 60 seconds. Pour off the iodine solution and gently wash with water. Shake off the excess

water from the surface. Decolorize with Gram´s decolorizer solution until the blue dye no longer flows

from the smear. Further delay will cause excess decolorization in the gram positive cells, and the purpose of staining will be defeated.

Gently wash the smear with water. Counterstain with Gram´s safranin solution for 60 seconds. Wash off the red safranin solution with water. Allow the slide to air-dry. Place 3 drops of Mounting Medium on the dried samples and cover it with a cover

slide Examine the slide under a microscope. Attention: Use the 100 x objective lens

(white ring) only after consulting the lab instructors (oil immersion objective). Attention! Wash off any spilled stain immediately with water to avoid leaving permanent marks in the sink, lab bench, or glassware. DISCUSSION After performing the staining, answer following questions: 2.4 Why do we heat fix the bacteria to the slide before staining?

A. So the bacteria hold the stain better. B. So the bacteria don’t wash off the slide. C. To kill them.

2.5 A basic dye has the color in the_____________ ion.

A. positive B. negative C. neutral

2.6 All bacteria have a slight negative charge; a basic dye has the color in the

positive ion. Opposite charges attract. This is the principle behind: A. Indirect staining. B. Why Gram-positive bacteria don’t stain pink. C. Direct staining.


2.7 Is this a direct stain or an indirect stain?

A. Direct stain B. Indirect stain

2.8 Is this a direct stain or an indirect stain?

A. Direct stain B. Indirect stain

II. MINIMUM INHIBITORY CONCENTRATION Answer these following questions before you join the practical course. 2.9 What does MIC stand for? 2.10 Calculate the concentration reduction (2) and the dilution concentration of each

tube in the serial dilution in Figure 11.

2.11 What is measured via a spectrophotometer?

initial sample concentration (mg/mL)

initial sample volume

Diluent volume

Total volume

Concentration reduction (2)

Dilution concentration (mg/mL)


INTRODUCTION In this section you will perform the antimicrobial agent susceptibility assay using dilution methods. The assay defines the minimum inhibitory concentration (MIC) which is the smallest amount of agent needed to inhibit the growth of a microorganism. You will be able to apply the knowledge acquired in previous Unit 1 regarding preparation of serial dilutions. To determine the MIC, a series of culture tubes is prepared and inoculated with the same number of microorganisms. Each tube contains culture medium with an increasing concentration of the agent. After incubation, the tubes are checked for visible growth (turbidity). The MIC is the lowest concentration of agent that completely inhibits the growth of the test organism. This procedure is also called the “tube dilution technique” (

Figure 10).

Figure 10. Antimicrobial agent susceptibility assay using the tube dilution technique to determine the minimum inhibitory concentration (MIC). Growth, which is assessed as turbidity, takes place in those tubes with antimicrobial agent concentrations below the MIC.

MATERIALS 15 mL culture tubes LB culture medium Antimicrobial agent Kanamycin Escherichia coli Set of micropipettes (P20, P200 and P1000) Beaker Set of tips Gloves


EXPERIMENTAL PROCEDURE (i) Antimicrobial agent susceptibility assay Perform following steps under a laminar flow hood: By using serial dilutions, prepare 4 culture tubes containing each 2.7 mL of LB*1

culture media with the antimicrobial agent Kanamycin*2 at following concentrations (in mg/mL): 5x10-1, 5x10-2, 5x10-3 and 5x10-4. Your stock solution is 5 mg/mL. Do not forget to label each culture tube (e.g. “[Kan] (5x10-1 mg/mL)”).

Figure 11. Serial dilution of Kanamycin*2 with a diluting factor of 10 for the Antimicrobial agent susceptibility assay. The stock solution (5 mg/mL) is placed in the very left tube and one part is mixed with nine parts of LB*1 culture media in another tube, which leads to a 1/10 reduction of the Kanamycin*2 concentration. Figure changed after: 1.5 mL tube, Test tube and Beaker: http://www.clker.com/cliparts/o/Z/8/K/a/X/eppi-lysat-b-hi.png, http://preview.turbosquid.com/Preview/2010/12/16__02_12_33/test%20tube_render.jpgff4d2eff-890e-47c6-ab72-da933fe249ecLarge.jpg, http://upload.wikimedia.org/wikipedia/commons/thumb/4/47/Beakers.svg/800px-Beakers.svg.png

Prepare an additional tube with 2.7 mL LB culture media without antimicrobial

agent. Do not forget to label the tube (e.g. “[Kan] (0 mg/mL)”) Add 10 µL of Escherichia coli suspension into each tube. Be sure that you shake

the bacterial suspension before taking the 10 µL aliquot. Take the 10 µL aliquot as fast as possible (before bacteria sediments) and inoculate it into the 2.7 mL culture medium.

Let the bacteria grow by shaking overnight @ 37 °C.

(ii) Measurement of bacterial growth by turbidity (spectrophotometry) Increased turbidity in a culture is an index of bacterial growth. A spectrophotometer does emit light and the amount of light transmitted through the sample is measured. The amount of transmitted light decreases as the number of cells in the sample


increases. Therefore the reading, called absorbance or optical density, indirectly reflects the number of bacteria. This method cannot distinguish between dead and living bacteria. An alternative method, called plate count method, reveals information related only to live bacteria. For more information on this latter method, please read Chapter 5, Brock Biology of Microorganisms, 13/e. Visualize the culture tubes and determine MIC. Determine MIC by measurement of bacterial growth via spectrophotometry as

follows: - Take 1 mL of LB culture media without Kanamycin and measure the optical

density (O.D.) at 600 nm. Set this value as blank. - Take 1 mL of LB culture media with 5 mg/mL Kanamycin and measure O.D. at

600 nm. - Take 1 mL of LB culture media with 5x10-1 mg/mL Kanamycin and measure

O.D. at 600 nm. - Take 1 mL of LB culture media with 5x10-2 mg/mL Kanamycin and measure



O.D. at 600 nm. LB*1 culture media: Lysogeny Broth (LB) or Luria-Bertani broth is the most widely used medium for the growth of bacteria. Formulation per one liter: 10 g Tryptone, 5 g Yeast Extract and 10 g NaCl. Antimicrobial agent Kanamycin*2: Kanamycin sulfate is a water-soluble antibiotic originally purified from the bacterium Streptomyces kanamyceticus. Kanamycin acts by binding to the 30S subunit of the bacterial ribosome and inhibiting protein synthesis in susceptible bacteria. DISCUSSION 2.12 Why do we discard 0.3 mL of the last tube of the Kanamycin dilution? 2.13 What do we measure in our sample with the help of the photometer, when we

set the LB culture media as blank? LITERATURE 1. Gregersen, T., Rapid method for distinction of gram-negative from gram-positive

bacteria, Eur. J. Appl. Microbiol. Biotechnol., 5, 123, 1978. 2. Madigan, M.T., Martinko, J.M., Stahl, D.A., and Clark, D.P., Brock Biology of

Microorganisms, Chapters 3, 5 and 26, ed. Benjamin Cummings, San Francisco, CA, 2012.

be cell microbio script.7-37

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