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Lab 1: Accurate Pipetting of Liquids
There are three main types of tools used in the lab to move liquids from one container to another. These
are: measuring pipets, Pasteur pipets, and the micropipettes (i.e. pipetman, piston pipette). This exercise
aims to get students used to dispensing liquids using these three different instruments.
Measuring Pipettes: Glass or plastic, calibrated to deliver any amount in the graduated scale from 1‐25
ml. Suction of liquid is through bulb or PipetAid. Serological pipettes are graduated to the tip meaning
that any liquid left in the tip after draining needs to be blown out. Volumes of 5, 10 and 25 ml‐pipets are
usually common in the lab. The 5 ml serological pipettes are accurate to ± 0.05 ml.
Pasteur pipette: A small, tapered glass tube, used with a bulb, and not graduated. It is used to dispense
liquid if the volume is not critical. Usually hold about 1 ml.
Micropipette: Used only with disposable pipette tips. This instrument is often used to dispense small
volumes ranging from 1 μl to 1000 μl. The three models that you will use are as follows:
 P20: 0.2 – 20 μl, calibrated in 0.02 μl increments.
 P200: 20 – 200 μl, calibrated in 0.2 μl increments.
 P1000: 200 – 1000 μl, calibrated in 2 μl increments.
Note: Do not use piston pipettes outside of their intended volume range! Why not? Two reasons:
1) the error rate is excessive and 2) you can break the pipette.
Pipetting techniques
Dispensing with mechanical air‐displacement pipettors demands skills and experience to do it right. There
are a few things not depending on the technique one should always pay attention to when pipetting:
* The pipettor/tip should be chosen to minimize the air space between the piston and the liquid.
* When filling the tip, the tip should not be placed too deep, but just under the surface of the liquid in the
reservoir (2‐3 mm).
* Pre‐wetting the tip improves both accuracy and precision.
* The pipettor should be held vertically, not at angle.
* The aspiration should be done smoothly, not too quickly.
* Dispense sample by touching the tip end against the sidewall of the receiving vessel to ensure complete
sample flow.
Factors affecting pipetting performance
The pipette tip
The tip is an integral component of the pipetting system and its shape, material properties and fit have a
considerable influence on the accuracy of liquid handling. In addition to fitting, most important is to test
how the tip wets, and whether there are droplets remaining after the sample is dispensed. To ensure
accurate pipetting results, only tips specified by the manufacturer should be used. Cheap, poorly fitting
tips not designed for the pipettor can result in serious measurement errors. If using tips other than those
specified by the manufacturer, one should always test the performance before beginning any experiments.
Especially the performance of filter tips used in wide variety of applications vary a lot depending on the
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pore size and material of the filter in addition to the properties of the tip. One should also keep in mind
that there is no such product as a universal tip.
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Environmental conditions
Sources of error from the environment include temperature (differences in temperature between the
pipettor, fluid and the ambient temperature), air pressure, and humidity. The single greatest contributor to
error is temperature, especially if working with air displacement pipettors (Joyce and Tyler, 1973; Lohner
et al.1996). As an example, increasing the temperature of the liquid from 5 °C to 28 °C while other
elements (pipettor and tip) are kept constant (22 °C), pipetting of 1 ml can have up to 6 % error in
volume. An ideal environment for pipetting maintains ambient temperature within 1 °C, including all
parts of the liquid handling system.
Inaccuracy and imprecision
Precision is an agreement between replicate measurements. Precision has no numerical value, it is
quantified by the imprecision. So high precision i.e. small imprecision, means very little variation
between the repeated measurements on the same sample. To achieve it you require a precision instrument,
but you must also follow good laboratory practice ‐ cleanliness and consistent correct handling. On the
other hand, it is possible to be very consistent, but consistently wrong. Inaccuracy is the numerical
difference between the mean of a set of replicate measurements and the true value ‐ so high accuracy i.e.
small inaccuracy means a very little difference between your mean sample and the true value. Accuracy is
achieved by careful calibration of a precision instrument. What is needed, of course, is both precision and
accuracy.
Formulas:
Expected value: what you set the pipette to measure
Measured value: what the balance says you measured
Accuracy (% error) = 100 x (Mean of measured value ‐ Expected value)/Expected value
Precision (% CV) = standard deviation/mean x 100
PART I. Practice Exercises
A. Pipetting with a PipetAid
First, we will practice dispensing different volumes of liquid using the 10 ml plastic disposable pipet. A
PipetAid is provided in each bench to make pipetting easy. This is an electric pipettor that replaces the
use of a manual aspirator. It has two buttons that control the flow of liquid in and out of the pipet. Each is
graded so that the farther you push the button, the faster the liquid is drawn up or expelled.
1. For each group, fill a 250‐ml flask halfway with water and obtain one 10 ml pipet.
2. Using this pipet, practice pipetting with the PipetAid. 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.
3. Learn how to control the rate of flow. Always have one eye on the pipet and at the same time
watch the level in the container.
4. Put less than 10 ml of water in the flask and then draw it all up without taking any air. If you do
pull air, measure the volume by the meniscus right below the layer of air bubbles.
5. Learn how to insert the pipet into a flask without touching the sides and hold it there while
withdrawing the liquid. This is a good sterile technique.
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6. Make sure you do not draw the liquid up to the cotton plug and especially not past the cotton
plug. This will contaminate your pipettor and requires disassembly and cleaning.
B. Using a Micropipette
A micropipette is designed to dispense small volumes of liquid. There are different types of micropipettes
designed for a particular range of volume. The volume capacity that each micropipette can dispense is
indicated at the top near the plunger. For practice, you will use three micropipettes of different volume
ranges.
Instructions:
1. Use three micropipettes: P20 (0.2-20 μl), P200 (20‐200 μl) and P1000 (200‐1000 μl) to cover a
range of volumes. There are two different sizes of tips‐ yellow or clear for the P20 or P200 and
blue for the P1000.
2. Put some water in a beaker to use to practice micropipetting.
3. Set a volume. Make sure that the volume you choose is within the range of the micropipette. Do
not push the volume dial beyond the indicated volume range, doing so will result in inaccurate
measurements and damage the instrument.
4. Load a tip onto the pipette by firmly pushing the barrel into the tip while it is still in the pipette
box. Tap the pipette up and down a few times to make sure it is seated properly.
5. Before you put the tip into the liquid, push the plunger down to the first stop. Do this outside of
your flask/tube so that you are not pushing potentially contaminated air into your potentially
sterile liquid.
6. Put the tip just below the surface of the liquid, then slowly release the plunger allowing the liquid
to be sucked up into the tip.
IMPORTANT: Never let the plunger snap back which may cause liquid to spurt into the piston
pipette and cause contamination and corrosion. If this happens, the pipette needs be cleaned.
Media or other liquids in the barrel may likely contamination your experiments.
7. After filling the tip, take the tip out of the liquid and make sure there are no bubbles or drops on
the outside.
8. Watch for leaky fits and air bubbles.
9. Place the tip against the edge of the tube/dish/liquid into which you are adding it and slowly
depress the plunger to expel the liquid. Push all the way to the first stop and then keep going to
the full stop to expel the liquid to its desired container.
10. Deposit the tip directly into the sharps container by pressing the ejector button.
11. Practice using the three micropipettes until you get used to them.
12. Never hold or place a pipette containing liquid into a horizontal orientation.
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PART II: Pipette Accuracy and Precision
A. Pipette tip volumes.
You will determine what volume each marking on the different varieties of lab pipette tips represents.
To do this, set the piston pipette to its middle volume and draw in water. With the pipette tip still
submerged in water, adjust the dial to increase or decrease the volume of water inside of the tip until
the meniscus lines up with the calibration mark being tested. Record the volume on your piston
pipette setting for every pipette tip marking.
Presentation of Data:
1. Draw each pipette tip for the P20, P200, and P1000 piston pipettes and their markings. Indicate
the volume each marking should represent and what the actual volume each pipette tip marking
is.
2. Turn in your drawings at the end of the lab period.
B. Piston pipette calibration.
Now you will determine whether your micropipettes are functioning correctly and you are using them
correctly. You do not want to assume they are correctly measuring volume; you want to be confident
that your experiments will be done correctly, so you want to make sure the pipettes are working
correctly. You will measure the accuracy and reproducible each of your piston pipettes. This will be
done by weighing the mass of water pipetted at various settings.
1. Accuracy - Pipette various amounts of water into a disposable weigh boat and determine the
volume of water you actually transferred knowing that the density of water is 0.998gram/ml (at
room temp); that is, 1 ml of water has a mass of 1 gram so 50μl (0.05ml) volume of water has a
mass of 0.05 gram (50mg).
2. Precision/Reproducibility - pipette the same volume multiple times. This will allow you to
determine the mean volume and see if you get the same reading each time. Try re‐using a tip
multiple times and see if that affects your measurements.
You will measure the mass of various volumes using each of the three piston pipettes and enter the value
of each measurement in the table below. Make sure you record all your data in your lab notebook as you
do the experiment (including the table below). This will allow you assess the accuracy of your
micropipettor and your technique at the beginning of the semester. It should not change if treated with
good care.
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Presentation of Data:
1. For this experiment, you each need to make several graphs that display your data; one graph each
from your measurements using the P20, P200 and P100 micropipettors. The X axis will be the
volume (in μl) and the Y axis will be the Mass (in mg or grams). Include error bars for each
measurement average data point.
2. Turn in your graphs to the instructor at the end of the lab period.
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Materials for this experiment:
1. Make sure all PipetAids in all the benches are working.
2. There should be a set of Micropipettes [P20, P200 and P1000] per group.
3. Analytical balance.
4. Plastic sterile 5 and 10 ml pipettes.
PART III: Serial Dilutions
It is sometimes necessary to determine the number of viable bacteria in a given culture. One way of doing
so is by inoculating an agar plate with an aliquot of a culture and counting the colonies after a 24 hr 37 °C
incubation period. However, even in a small amount of culture media, for example 100 μl, there may be
billions of bacteria. Plating an undiluted sample may result in a lawn of bacteria making it impossible to
count colonies or to select an individual colony if desired. Therefore, cultures often need to be diluted at
least 1 million fold or more before plating to be able to observe individual colonies. Diluting to such a high
dilution fold would mean that one needs to dilute 1 ml of culture into 1000 L of media. It’s unlikely that
one has a 1000 L container to make such a dilution, and it would be economically prohibitive to make such
a single dilution. Instead, a serial dilution is used to dilute the culture several times to reach the desired
dilution factor. For example, to make a 1 million fold dilution, one would make six individual 10 fold
dilutions in series. To perform such a serial dilution, 100 μl of culture is added to 900 μl of media to make
the first 10 fold dilution. Then 100 μl of the 10 fold dilution is added to another 900 μl of media to make
the 100 fold dilution, and so on, until the original culture has been diluted up to 1 million fold. The culture
from each dilution is then spread onto agar plates which are then incubated overnight at 37 °C. Each
individual bacteria will grow into an individual colony known as a colony forming unit (CFU). The number
of bacteria in the original culture can be calculated by multiplying the final number of colonies on the plate
by the dilution factor.
In this activity you will quantify the CFU of Escherichia coli BL21 λDE3 pET3a-4 using the serial dilution
and plate count method.
Safety reminder: Handle bacterial cultures carefully, for example while opening microfuge tubes to prevent
aerosolization. Use aseptic technique and wear appropriate PPE (i.e. gloves, lab coat, and goggles).
Instructions:
1. Label seven sterile microfuge tubes with the dilution factor (10, 102, 103, 104, 105, 106, and 107).
2. Label seven LB-agar plates with the dilution factor (10, 102, 103, 104, 105, 106, and 107).
3. Using aseptic technique, pipette 900 μl of LB broth into each microfuge tube.
4. Using aseptic technique, pipette 100 μl of Escherichia coli BL21 λDE3 pET3a-4 culture into the
microcentrifuge tube labeled “10”. Mix by vortexing or pipetting up and down at least 5 times.
5. Remove 100 μl of the diluted cell culture from the tube labeled 10 and add it to the tube labeled
102. Mix by vortexing or pipetting up and down at least 5 times.
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6. Remove 100 μl of the diluted cell culture from the tube labeled 102 and add it to the tube labeled
103. Mix by vortexing or pipetting up and down at least 5 times.
7. Continue to serially dilute the bacterial culture into each consecutive tube until the tube labeled
107 is reached.
8. Mix the diluted cell culture in the tube labeled 10 and using aseptic technique, pipette 100 μl onto
the LB-amp agar plate labeled 10.
9. Sterilize an inoculating loop or bacterial spreader by flaming, and then allow it to cool.
Important: DO NOT use a flame on disposable plastic spreaders, they are intended for single use.
Very gently spread the bacterial culture over the entire surface in all direction including the edge
of the plate. Do not us the streak plate technique – you are not isolating single colonies.
10. Plate 100 μl of each dilution on the matching plate using the instructions provided in step 9.
11. Stack the plates upside down and place in the incubator at 37 °C for 16-24 hrs.
Results and Presentation of Data:
Retrieve your culture plates from the incubator and arrange them on your bench from least to most dilute.
Record the number of CFU across all plates. Plates that have bacterial growth so dense that no single
colonies can be seen should be recorded as having a “lawn” of bacteria. Plates having more than 200
CFU should be recorded as “too many to count”.
To count colonies, use a sharpie marker to place a dot on the bottom of the plate in the middle of each
colony. Count each plate that has between 30-200 CFUs, and multiply the number of CFU by the dilution
factor for that plate. Remember, you also need to take into consideration the fraction of the dilution that
was inoculated, since only 100 μl of the 1,000 μl was spread on the plate (that’s a factor of 10 fold since
the CFU would have been 10 fold higher if the entire 1000 μl was plated).
Concentration of bacteria (CFU/ml) = # of CFU x dilution factor x
1,000 μl
100 μl
1. Record the number of CFU across all plates.
2. Calculate and report the concentration of bacteria per ml (CFU/ml) in the original culture tube
using the results from each individual plate.
3. Then average the calculated original concentration values from the all of the plates that have 30200 colonies and report the value to the nearest significant figure and include a standard
deviation.
4. Did each serial dilution sample have exactly 10 times the number of CFU as the previous
dilution? If not, how different were the samples and why were they not exactly proportional?
5. How could you change the experiment to more accurately determine the number of CFU in the
original culture?
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Part IV – Making LB Media and LB-Amp Agar Plates
Bacterial cultures are grown in growth media. A common growth media for growing E. coli is Lysogeny
Broth (LB) (also referred to as Luria-Bertani broth). LB is used as a liquid media, and can also be
formulated with agar to make plates. There are various recipe modifications to LB and LB agar when
there is a desire to tailor a media for a particular experiment.
Standard liquid LB is composed of water, tryptone (a tryptic digest of casein (milk protein) supplying an
assortment of peptides), yeast extract (to supply vitamins and trace elements), and NaCl (to provide
osmotic balance). Pre-mixed dry LB may be purchased from various manufacturers. LB agar has a
similar as liquid LB with the exception that agar is included in the formulation. Agar is composed of a
mixture of the polysaccharides agaropectin which are obtained from algae where they function as
components of the cell wall. Because of its gelling properties, agar is used as a substrate to contain
culture media and is added to liquid media for solidification (similar to gelatin).
In this activity you will make liquid LB broth as well as LB agar plates with the antibiotic ampicillin.
Instructions:
A. LB Broth:
1. Label a 100 ml bottle or flask with the name of the medium, date of preparation, and group
initials.
2. The following is a recipe to make 1 L of LB broth (and LB-agar):
Add the following to 800 ml H2O
 10g Bacto-tryptone
 5g yeast extract
 10g NaCl
 Optional: Add 18 g of agar if making LB-agar
 Adjust pH to 7.5 with NaOH
 Adjust volume to 1L with dH2O
 Sterilize by autoclaving
3. You will make 50 ml of LB broth. Calculate how much of each ingredient you will need to make
50 ml. Create a table in your lab notebook and follow your recipe measurements.
For 50 ml, add the following to ______ ml H2O
 _______ Bacto-tryptone
 _______ yeast extract
 ________ NaCl
 Optional: Add ______ g of agar if making LB-agar
 Adjust pH to 7.5 with NaOH
 Adjust volume to 50 ml with dH2O
4. Show your calculations to your instructor before proceeding,
5. Weigh out and dissolve all of the ingredients for LB broth in a beaker or flask.
6. Transfer liquid to a bottle and cap very loosely. Place a small strip of autoclave tape on top of the
cap. Hand over the bottle to your instructor for autoclaving.
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7. After autoclaving the LB broth will be good to use for a year.
B. LB-Amp Agar:
1. Label a 250 ml flask with the name of the media, date of preparation, and group initials.
2. Weigh out and dissolve all of the ingredients for LB-agar in a 100 ml flask.
3. Cover the flask with aluminum or cap very loosely and place a small strip of autoclave tape on
top of the cap or aluminum.
4. Hand over the bottle to your instructor for autoclaving.
5. During the autoclaving procedure, label two petri plates with the name of the medium, date of
preparation, and group initials.
6. After autoclaving, incubate the autoclaved LB-agar in a 55 °C water bath to allow the LB-agar to
cool. Wait approximately 15 min for the LB-broth to cool (longer for larger volumes).
7. Once the LB-agar has cooled, add ampicillin. A 1000X stock of ampicillin has a concentration of
100 mg/ml (dissolved in water). How much 1000X stock will you add to the 50 ml LB-agar?
Hint: add 50 μl of the 1000X ampicillin stock.
8. Gently swirl the flask without causing air bubbles to form.
9. Pour approximately 20 ml of the LB-amp agar into 2-3 separate 60 mm petri dishes. Keep the lid
over the plates to prevent
10. Let the LB-agar solidify. Once solidified, turn the plates upside down and store at 4 °C. The LBamp plates will be good for 2 months.
11. Post lab questions:
 Why is it necessary to cool the agar prior to adding ampicillin?
 Why is autoclave tape used?