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Introduction to BIO 151 (Week 1)
This handout contains basic information and skills that you will need to
successfully design, execute, and complete an independent research project
in forensic biology this semester. Because this is an advanced inquiry-based
course, students will be responsible for all aspects of their research projects,
including experimental design, preparing their own materials and reagents,
choosing the appropriate lab water and mixing tools, disposing of
biohazardous and chemical waste, and cleaning up at the end of each lab day.
We will discuss the material in this handout on the first day of class and you will be given a homework
assignment to help you practice and hone your skills and understanding. It is vital that you pay close
attention to the information below so that you can work safely, efficiently, and effectively in the lab.
I.

Safety and Cost Considerations
Gloves. Gloves should be worn at all times. To save money, please change your gloves sparingly
and think critically about whether or not changing gloves is actually necessary. Unless you have
an allergy to latex, please use the latex gloves. Nitrile gloves are more expensive.

Bench Diapers. Always work on a piece of bench diaper. Change the diaper if it gets wet or dirty
or if keeping the same piece of bench paper in place may lead to cross-contamination of evidence.
Think critically about whether or not it is necessary to change your bench diaper.

Cleaning Scissors and Tweezers. As in BIO 150, you will be using three solutions to clean your
scissors and tweezers between uses: 10% bleach (to destroy DNA), dI water (to remove the
bleach), and ethanol (to destroy any bacteria or other microorganisms). Bleach is fairly effective
at preventing cross-contamination of DNA from one evidence item to another, but is not full-proof.
You may wish to clean your non-disposable instruments more than once between uses to make
sure to avoid inadvertently transferring minute amounts of DNA from one
item to another. In addition, you should always perform your negative
controls first, your experimental samples second, and your positive controls
last. You can save your bleach and water beakers by covering them with
seran wrap and placing them in your lab cabinet between labs. However, the
beakers containing ethanol should be emptied into the ethanol container in
the chemical hood for storage in a flame proof cabinet between uses.

Autoclaving. Think carefully about whether or not items need to be autoclaved. Sometimes it is
not necessary to sterilize instruments and reagents, while other times it is very important. Look at
the labels on reagents and items. Do the items come pre-sterilized (e.g. filter tips, disposable
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scalpels)? If so, sterilizing them is a waste of time and resources. In addition, although you need
to clean your non-disposable implements (scissors, tweezers), you do not need to sterilize them
between uses (except by briefly cleaning them with ethanol). Likewise, you can clean beakers and
other types of glassware with soap and tap water; in most instances, you do not need to sterilize
these items prior to use. Your decision on whether or not to autoclave items will depend on
whether contamination with bacteria is likely to interfere with your downstream analyses, whether
the item you are examining is already compromised with bacteria, and whether the reagent is
subject to bacterial contamination. For example, you do not need to autoclave ethanol since
bacteria cannot grow in it! Try to think critically and logically about sterilization, and never be
reluctant to ask questions if you are confused. Remember that sterilizing items is time-consuming
and expensive. More information on autoclaving is provided in Section IV of this handout.

Lab water. Review Section V of this handout as a guide to choosing the right type of lab water for
the task at hand. Know the differences between tap, deionized, distilled, and nuclease-free waters
and when it is important for water to be sterile. Highly purified waters are
critical for some applications but are very expensive are not necessary for
most applications. For example, rinsing glassware in nuclease-free water
would quickly bankrupt a lab, but failing to use nuclease water when
setting up a PCR reaction could result in DNA degradation and the loss of
probative genetic information from a sample. Again, don’t hesitate to ask
questions if you are confused and need some guidance on which type of
water to use.

Biohazardous and Chemical Waste. Biohazardous waste is any organism, or its toxins, that can
cause disease in animals or humans. Biohazardous waste is produced by living organisms and can
cause damage to other biological organisms. Most biohazardous waste can be
neutralized with bleach or by autoclaving. In contrast, Chemical waste is waste
made from harmful chemicals. It is non-living and usually cannot be
neutralized by bleach or autoclaving (either of which might actually do more
harm than good). Chemical waste should therefore always be collected in
separate collection vessels from biohazardous waste. In addition, each type of
chemical waste is subject to different neutralization procedures and so must be
collected separately from other types of chemical waste. If an item has both
biohazardous and chemical waste properties, the neutralization of its
biohazardous component takes precedence. However, since the item also contains hazardous
chemicals, the method of rendering it free of biohazardous materials must be chosen very
carefully. In lab, we will have several, clearly labeled containers for collection waste. Be sure to
use the right collection vessel and ask if you need help.

Material Safety Data Sheets (MSDS) are available for almost every chemical, and companies
that make reagents with proprietary formulae must disclose the chemical and/or biohazardous
properties of their products. MSDS information can be used to determine whether a chemical or
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solution is biohazardous or contains chemical waste and how to dispose of it appropriately.
Fortunately, in most laboratories (including this one) waste disposal is delegated to specialized
personnel, who are an excellent source of information and help to researchers and technicians.
These specialists supply labs with labels, collection vessels, and waste protocols that meet
government guidelines, and run central facilities where labeled waste can be deposited. MSDS
access will be provided; however, you will be given clear directions on where to dispose of
materials, so you probably will not need to access the MSDS materials while you are taking this
course.
II.
Calculations and Measurements
Performing accurate calculations and measurements is absolutely
critical in a molecular biology laboratory. To succeed, you will need to
be fluent in unit conversions, know which equations to apply for which
types of calculations, know the capabilities and limitations of your
equipment, and be able to maintain your equipment for optimal
performance. For example, micropipetters have set ranges and should
not be used outside of those ranges. In addition, they should be calibrated regularly (about once every
6-12 months is optimal).
Homework 1 is designed to help you test your current skills and, if they are deficient, to become
comfortable solving routine types of lab calculations. Remember: you are striving for fluency, which
means that you should be able to set up and solve problems easily and automatically. You should also
be able to quickly identify what additional information (if any) you will need to solve the problem and
how to get it.
1. Measuring Volumes of 1 uL - 1 mL: Use and Maintenance of Micropipetters
Micropipetters are indispensable to molecular biologists because they allow the accurate measurement
of volumes between 1uL and 1 mL - the typical volume range used for most molecular biology lab
work. If you have not done so already, it is very important that you commit to memory the following
simple units of the metric system of measurement. You will use them often!
“milli” (m)= 10-3
“micro”) (µ) = 10-6
“nano” (n) = 10-9
“pico” (p) = 10-12
For example, 650 mL = 0.65 L and 200 uL = 0.2 mL. If these types of conversions are not second
nature (and very easy to do in your head), start practicing!
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The micropipetters we will use are called
Finnpipettes (Fisher Scientific). Each instrument
is calibrated for a narrow range (0.5-10 µL, 5-50
µL, 20-200 uL, and 100-1000 µL) and is a
different color. The 0.5-10 uL micropipetter is
used with microtips when barrier filters are
needed (e.g. for DNA work) but can be used with minitips for most other purposes. The 5-50 uL and
20-200 uL pipettes are used exclusively with minitips and the 100-1000 uL pipette is used with
macrotips. Fortunately, if you try to put the wrong size tip on a micropipetter, it will not fit.
Unfortunately, it is easy to confuse 20 uL with 200 uL (etc.) if you are not familiar with the
instruments. Everyday use will limit such mistakes; however, it is important to be very careful!
Fill in the far right column of the table below. If you have questions, ask!
READING THE VOLUME WINDOWS ON THE FINNPIPETTES
READING IN VOLUME
SAMPLE
VOLUME OF SAMPLE READING
WINDOW AT TOP VALUE
READING
(FILL IN)
200-1000 uL
1-0-0
0-7-7
20-200
50-200 uL
2-0-0
0-7-7
5-50
5-50 uL
5-0-0
0-7-7
0.5-10
0.5-10 uL
5-0-0
0-7-7
PIPETTER
RANGE
100-1000
To suck liquid into the tip: Gently press down on the plunger to the first stop position (the
position where you will first feel resistance to your pressing.) Then place the tip into the liquid to be
measured (below the level of the liquid’s surface) and gently and slowly release the pressure on the
plunger. Then remove the pipette from the liquid and transfer the contents in the tip to the container
you wish to fill. (The liquid will remain in the tip as you do this because it is being held in by the
vacuum you created inside the micropipetter when you released the pressure on the plunger.)
To expel the liquid from the tip: Gently push the plunger down to the first stop position and then
continue until you reach the second (final) stop position. You may use either your thumb or your first
finger to depress the plunger, whichever is most comfortable. If you are transferring the contents of
the tip into a tube that already contains liquid, gently express the contents into the liquid, below the
level of its surface. If you are transferring the contents of the tip into an empty tube, place the tip on
one of the inside surfaces of the tube while expelling. (This will allow capillary action to help you
release all the contents of the tip into the tube; if you do not do this, some liquid may remain in the
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tip.) As you expel, keep the pressure on the plunger. Do not release the pressure until you
have removed the tip from the tube. (Otherwise you’ll just suck the liquid right back up into the
tip!)
The best technique is to watch the liquid as you are micropipetting to be absolutely sure that you get
the correct amount in the pipette tip at the start of the process (no bubbles) and that all of it was
expelled at the end. Experienced lab technicians develop a good sense of how different volumes
appear in a tip. This helps them avoid pipetting errors, particularly those that occur when they
accidentally pick up the wrong pipetter.
The micropipetters we will be using have an ejector on one side. After use, the tip can be popped off
by pushing down on the ejector button at the top of the pipetter. This avoids the necessity of having
to remove the pipette tip with your fingers.
2. Weighing Reagents
To accurately measure the weight of a dry substance, particularly when the amount is small, plastic
weigh boats or weigh papers are a good choice. A small spatula is used to carefully transfer the
powdered reagent to the weigh boat or paper and the substance is then weighed on a scale. Since the
weight of the boat or paper needs to be subtracted from the total weight in order to get an accurate
measurement, the weigh boat or paper is first placed on the scale and the scale is "zeroed" or "tared"
(set to 0) before the substance is added for weighing. Scales have different levels of sensitivity and it
is important to know the limits of the scale you are using. This is usually displayed on the back or side
of the scale.
When making solutions in the lab, it is critical that you know the difference between weight, moles,
molecular weight, and molarity. These terms are defined below. Make sure you commit them to
memory if you don’t already know them.
Weight is a measurement of the gravitational force acting on an object. Near the surface of the Earth,
the acceleration due to gravity is approximately constant; this means that an object's weight is
roughly proportional to its mass wherever the measurement is made (e.g. Sacramento or Zanzibar).
Therefore, because all our measurements will be made on planet Earth (there will be no field trips to
Venus or Mars this quarter!) we will use the terms weight and mass interchangeably. Please
remember, however, that weight and mass are not equivalent terms in physics.
A mole is defined as 6.022 ×1023 atoms or molecules of a substance. This may seem like a
ridiculously high number, but given the small size of most molecules used in molecular biology, it is
actually very useful.
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Molecular weight (MW) or formula weight (FW) is defined as the weight of one mole of an atom
or molecule. The most common unit of expression for molecular weight is g/mol. NaCl is a very small
molecule, so 6.022 ×1023 molecules (one mole) of NaCl doesn’t weigh much (58.44 grams or about 2
ounces). On the other hand, one mole of human chromosome 1 (a molecule containing 247 million
base-pairs of DNA, with a MW of 660 g/mol per DNA base-pair) weighs a lot more. Use the space
below to calculate the MW of chromosome 1.
Assuming that there are 6 billion human beings on Earth and each human body contains about 10
trillion cells, how many moles of human chromosome 1 are on the planet?
Taken together, how much do all chromosome 1’s on the planet weigh? Calculate the number in
grams, pounds, and tons. (Note: There are 453.6 g per pound and 2,000 pounds per ton.)
To give you some perspective, the U.S.S. Missouri (a typical battleship) weighs about 45,000 tons.
Molarity (M) is defined as the number of moles per liter of a solution of a substance. Obviously, the
molarity of a solution can be manipulated by a researcher since it depends on how much of the
substance is added to how much solvent (usually water). For example, to make 1 liter of a 1 M
solution of NaCl would require 58.44 g of NaCl. To make 1 L of a 0.1 M solution, you would need only
5.844 g. (How many grams of chromosome 1 would you need to make up 1 L of a 1M solution?)
3. Working with Solutions
The concentrations of solutions in the lab are usually given in molarity (M = moles/L). However, they
may also be expressed in grams/L, as a percent, or as an “X” (e.g. 10X, 1X, etc.). By definition, 1
g/mL = 100%, so conversions can easily be made between the first three methods of expressing
solution concentration (assuming that you know the FW of the substance). “1X” is usually the working
concentration of a solution. Without additional information, you can’t convert “X” to M, g/L, or a
percent. However, if your stock solution is at 10X, you can easily make a 1X solution by diluting the
solution 10-fold.
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Here’s a practice problem: NaCl has a FW of 58.44. You have a 5 M solution of NaCl. What is its
concentration in g/L? What percent solution is it?
(5 moles/L)(58.44 g/mole) = 292.2 g/L
(292.2 g/L)(100%/(1 g/mL)(1 L/1000 mL) = 29.2%
You will need to learn to do these kinds of conversions quickly and easily. Make a habit of always
including units and cancelling them out (no shortcuts!). You will catch errors much more easily if you
do this. It also helps to identify first what units you want for your final answer. This unit should be in
the numerator in one of the fractions in your conversion.
For example: How many moles are in 3.0 mL of a 10.0 g/mL solution of NaCl? You want moles in your
answer, so moles should be in the numerator of one of the fractions in your calculation.
(1 mole/58.44 g)(10 g/mL)(3 mL) = 0.51 moles
In addition, you will find the following formula very helpful when working with solutions in the lab:
(Vi)(Ci) = (Vf)(Cf)
Where: Vi = initial volume
Ci = initial concentration
Vf = final volume
Cf= final concentration
Suppose, for example, that you have a stock solution of EDTA at 2 M. You wish to make 50 mL of a
0.8 M solution of this reagent. How would you do it?
Vi = (Vf)(Cf) = (50 mL)(0.8 M)
Ci
= 20 mL
2M
Assuming that the solvent is dI water (in molecular biology it usually is), you would add 20 mL of 2M
EDTA to 30 mL of dI water to make 50 mL of 0.8 M EDTA solution.
III. Lab Water
For the purposes of this class, we will divide our water supply into the two types usually available in
molecular biology laboratories: tap and purified. Each can then be sub-divided into classes, as shown
in the table.
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Type
Method of Purification
Properties
Tap
Industrial
None
Contains ions, organic compounds (natural and manmade), particulates (vegetable debris, sand, and
colloids), microorganisms (>1 cfu/mL), DNAases, and
RNAases; Free of microorgansims and DNAases if
sterilized
Drinking
Gross filtration (sand,
activated charcoal) and
disinfection (chlorine,
UV)
Contains ions, organic compounds (natural and manmade), microorganisms (< 1 cfu/mL), DNAases and
RNAases; Free of microorganisms and DNAases if
sterilized
Ion exchange resin or
electrodeionization
Very low levels of mineral ions; contains organic
compounds, microorganisms (<1 cfu/mL), DNAases
and RNAases; Free of microorganisms and DNAases if
sterilized
Distilled
Distillation
Very low levels of ions and organic molecules;
microorganisms may also be present at low levels,
DNAases and RNAases. Free of microorganisms and
DNAases if sterilized
Ultrapure
Ultrafltration; reverse
osmosis
Free of ions, organic molecules and microorganisms
but not necessarily DNAases and RNAases; DNAases
can be removed by sterilizing
Nuclease-free
Ultrafiltration; reverse
osmosis; DEPC
treatment
Free of ions, organic molecules, bacteria, DNAases,
and RNAases.
Purified
dI (deionized)
Industrial Water can be used for washing glassware, plastic ware, hands, etc., but it is not safe to
drink. It usually comes directly from a tap located in a lab sink and is often labeled as “Industrial
water” or as “Unsafe to drink.” Once glassware or plastic ware has been cleaned with industrial water,
it should always be rinsed thoroughly with dI water to remove any residual contaminants.
Deionized (dI) water has largely replaced distilled water as a source of ion-free water. It is less
expensive to produce than distilled water and, since most of the chemical contaminants in water are
ions, removes the chemicals of most concern. dI water can be sterilized by autoclaving and is suitable
for the preparation of most molecular biology lab solutions and for rinsing glassware and plastic ware
after it has been washed in industrial water. Because it is does not contain minerals that leave
deposits, it is also the best choice for filling water baths and piping into autoclaves and lab
dishwashers. It is also used to prepare microbiological media prior to autoclaving (although sometimes
it is better to use tap water because some microbes require ions for robust growth).
Ultrapure water is made by filtering dI or distilled water (in a reverse osmosis system) through a
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very fine filter (a pore size of 0.22 um is typical) to remove any residual ions and bacteria. However,
DNAases or RNAases may still be present. DNAases can be removed by autoclaving since they are
permanently denatured by exposure to heat. Autoclaved ultrapure water is the best choice for the
preparation of any solution that will come into contact with DNA, including solutions for DNA
extraction, quantitation, and PCR. However, it is not sufficiently pure for experimental solutions that
require intact RNA (such as reverse transcriptase PCR).
Nuclease-free water is ultrapure water that has been treated with DEPC (a non-specific inhibitor of
RNAases) and then autoclaved (to destroy the DEPC and inactivate DNAases). RNAases are not
destroyed by autoclaving because they tend to refold after being heat-denatured. Nuclease-free water
is necessary for any lab work that requires intact RNA.
IV.
Autoclaving
An autoclave is an essential piece of equipment in a molecular biology lab.
It is used to sterilize solutions, supplies, small instruments, and
biohazardous waste using high pressure steam (at 120 deg C or above).
Steam is used because it can achieve sterilization much more quickly than
hot air (15 minutes as opposed to several hours).
Not all materials can survive the high temperatures and pressures
produced in an autoclave cycle and it is therefore extremely important to
use the right type of glassware and plastics. Thick Pyrex glassware and
either PFA, polypropylene, polysulfone and Noryl plastics are usually
autoclave-safe, but you should always check to make sure that the items
you place in the autoclave can survive the procedure. Thin glass will break, and many types of plastics
will melt, leaving a big mess and producing volatile products that may be toxic.
Extreme care should be used when loading and unloading an autoclave. It is very easy to get burned –
either by inadvertently touching the side of the autoclave with your uncovered arm or by failing to
move out of the path of the steam as it escapes from the autoclave door. Long protective autoclave
gloves should be worn at all times, and the door should be opened in such a way that your body is out
of the path of the steam. It is always best to place items in an autoclave-proof tray during the run so
that if breakage occurs it is contained. Trays also make removing items from the autoclave safer and
easier.
Autoclave tape changes color from white to striped brown during autoclaving. Placing autoclave tape
on an item prior to autoclaving provides you a visual cue that the item or solution has been sterilized.
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During autoclaving, some water will evaporate from your solutions. Therefore, to maintain the right
concentration, you should use a Sharpie to mark the level of the fluid before autoclaving. After
autoclaving, you should then add STERILE water to the solution to replace the water that was lost
during the procedure. If you do not do this, your solution may be more concentrated than you
intended, since only the water evaporates during the process (not the ions or other solutes).
V. Mixing
During molecular biology experiments, you will often need to mix two or more liquid ingredients
together. Mixing is extremely important, and experiments can fail if you do not mix ingredients
together thoroughly. However, it is important to know the various methods of mixing and when to
apply them. Vortexing is fine under many circumstances but can be destructive in others. Usually, the
protocol you are following will indicate what method of mixing is appropriate. If not, follow the general
rule that solutions containing DNA, RNA, and proteins that are needed for downstream applications
should be mixed using the flicking or pipette method. Most other mixtures are amenable to vortexing
unless they contain detergents (e.g. lysis buffers) that will become bubbly when vortexed.
1. Vortexing
Most molecular biology vortexers have 3 settings: Off, Continuous, and Touch. The Touch setting is
preferred because the vortexer will not operate (and make a lot of noise) unless the researcher places
the tube in the rubber holder and applies downward pressure. In the Continuous mode, the vortexer is
on all the time, regardless of whether anything is being vortexed. Some vortexers have tube
attachments that allow researchers to shake contents vigorously over an extended period of time.
2. Flicking, Pipetting, and Inverting
The materials in tubes can also be mixed by gently flicking the tube several times with your finger.
Alternatively, the contents can be gently mixed by pipetting the mixture up and down a few times. It
is usually important not to introduce bubbles, so it must be done carefully and gently to be effective.
Another method is to simply invert the tube several times, making sure the lid is firmly in place.
VI.
Supplies Boxes
Each student will be assigned his/her own supplies box for use during the semester. The materials in
your box are expensive and the box must always be locked in your lab cabinet between labs. Each
box contains the following items, and you are responsible for making sure that nothing gets lost!
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
6” plastic ruler

Set of four micropipetters (Finnpipettes) and rack

One microcentrifuge tube opener

Reversible rack for 1.5-mL/0.5 mL microcentrifuge tubes

Rack for 0.2-mL PCR tubes

Roll of lab tape (colored)

Roll of autoclave tape

Roll of evidence sealing tape

Black ultrafine Sharpie marker

Peel-off China marker

Pair of tweezers

Pair of scissors

Lab timer

1 box small Whatman filter paper circles

1 dropper sterile, distilled water

1 pack of lens paper

1 magnifying glass
During the first day of lab, you will be asked to inventory your box to make sure everything is present.
You will then sign the inventory form and submit it to your instructor.
VII.
Tips, Tubes, and other Items
Before beginning your lab work, you and your lab partner will need to gather together the following
items, which you can store in your lab cabinet/drawer:

One box Kim Wipes

One 100-mL graduated cylinder

One freezer box

Two plastic collection beakers labeled “dry waste” and “liquid waste

2 full pipette tip racks (1 macro, 1 mini)*

One large beaker filled with 1.5-mL tubes*

One medium beaker filled with 0.5 mL tubes*

Three 250-mL autoclave-ready bottles

One 500-mL autoclave-ready bottle half-filled with dI water*
Use the colored tape in your supplies box to label the items that are not in your supplies box with the
bench numbers of you and your lab partner (e.g. 1/8). Use the Sharpie in your supplies box to label
the tape; DO NOT label directly on the items with the Sharpie.
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The items marked with an asterisk will need to be sterilized immediately. Follow the instructions below
to autoclave these items:

Place a piece of autoclave tape on the side and top of your tips boxes. Put a piece of
lab tape on the side of each box and label it with your lab station number. Place the
tip boxes on the autoclave cart.*

Fill your large glass beaker with 1.5-mL microcentrifuge tubes and your medium glass
beaker with 0.5 mL microcentrifuge tubes. Cover each with aluminum foil and adhere
a small piece of autoclave tape to the top of the foil as well as on the side of the
beaker. Label the beakers with your lab bench numbers (using your colored lab tape)
and place the beakers on the autoclave cart when you are done.

Add approximately 400 mL dI water to your 500-mL autoclave bottle. (Your instructor
or T.A. will show you where the dI water taps are located.) Label the bottle with the
name of the solution (“Sterile dI water”), your lab bench numbers, and today’s date.
Place autoclave tape on the cap and side of the bottle and then place the bottle on
the autoclave cart. Be sure to loosen the lid to prevent the bottle from breaking
during autoclaving.

Prepare your solutions of Tris and EDTA (see below).

When all the items are ready, your instructor or T.A. will instruct you in the proper
use of the autoclave.

When your items are finished autoclaving, they will be removed from the autoclave
and allowed to cool until the next lab period. When they are completely cool, you can
store them in your lab cabinet.
*The tips in these boxes can be used for most applications. However, you will be provided with
pre-sterilized, nuclease-free, aerosol-barrier tips for your qPCR work.
VIII.
Prepare T10E0.1 Buffer
You will eventually need TE buffer for qPCR. To practice making up a solution, you will prepare your TE
buffer during the first week of lab, following the steps below.
1.
Prepare 100 mLs of 1 M Tris-HCl, pH 8.0.

The poly bottle of Tris Base is on the front bench.

Using the FW on the bottle, calculate how much powdered Tris Base you will need.

Weigh out the appropriate amount of Tris Base and place it in a 250-mL beaker

Add 70 mLs dI water and 4.2 mL concentrated HCl. Add a stir bar, place the
beaker on a stir plate, and stir until dissolved.
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
When the solution is clear, test the pH. Make minor adjustments to bring the pH to
8.0 using either concentrated HCl or NaOH crystals.

Pour the solution into a 100 mL graduated cylinder and top off to exactly 100 mLs
with dI water.

Pour the solution into a 250-mL autoclave-ready bottle and mark the miniscus with
your Sharpie.

Label the solution “1 M Tris-HCl, pH 8.0” followed by today’s date and your lab
bench numbers.

Place a small piece of autoclave tape on the lid and side of the bottle, crack the lid,
and place the bottle on the autoclave cart.
2.
Prepare 100 mLs of 500 mM EDTA, pH 8.0

The poly bottle of EDTA is on the front bench.

Using the FW on the bottle, calculate how much powdered EDTA you will need.

Weigh out the appropriate amount of EDTA and place it in a 250-mL beaker.

Add 80 mL dI water

Add 2g NaOH pellets

Add a stir bar, place the beaker on a stir plate, and stir until dissolved. (This may
take a little while.)

After the solution is clear, test the pH. Make minor adjustments to bring the pH to
8.0 using either concentrated HCl or NaOH crystals.

Pour the solution into a 100 mL graduated cylinder and top off to exactly 100 mLs
with dI water.

Pour the solution into a 250-mL autoclave-ready bottle and mark the miniscus with
your Sharpie.

Label the solution “0.5 M EDTA, pH 8.0” followed by today’s date and your lab
bench numbers.

Place a small piece of autoclave tape on the lid and side of the bottle, crack the lid,
and place the bottle on the autoclave cart.
3.
After your autoclaved solutions have completely cooled, prepare a 5 mM solution of EDTA as
follows:

Use a micropipetter to add 9.9 mL sterile water to a 15-mL sterile Falcon tube.
Add 0.1 mL of your 0.5 M EDTA solution. Invert to mix. Label the tube “5 mM
EDTA, pH 8.0” followed by today’s date and your lab bench numbers.

Calculate how much 1 M Tris (8.0) and 5 mM EDTA (8.0) you will need to make 10
mL of a solution that contains a final concentration of 10 mM Tris and 0.1 mM
EDTA. Write out the recipe and check with your instructor or T.A. before
proceding:
FOR 151
Fall 2010
Page 13
Version: 2/18/2016
T10E0.1 Buffer Recipe (10 mL):
___________ mL 1M Tris (8.0)
___________ mL 0.05 M EDTA (8.0)
___________ mL sterile water

Use micropipetters to mix together the ingredients in a sterile 15-mL Falcon tube.
Label the tube “T10E0.1 Buffer” followed the date and your lab bench numbers.
All your solutions should be stored in your lab cabinet for future use.
FOR 151
Fall 2010
Page 14