Objective
InnovaBio positively impacts the community by giving students realworld research experience working on projects for life science
companies in need of innovative technical solutions for research
problems. InnovaBio offers scientific expertise and a low-risk financial
environment that benefit both the partner enterprise and the students
participating in the scientific work. Our objective is to mentor the next
generation of Biotechnology Graduates and simultaneously assist local
scientific and biotech enterprises in developing research projects.
To assist yourself and your team in fulfilling the objective, please
do the following steps:
Step 1 : Be Aware of your experimental plan: know what it is
and where to find protocols.
Step 2: Understand the steps within the protocols or
intended experiments, and how the experiments fit into the
overall project goals.
Step 3: Be confident in what you are about to embark on is the
correct experiment for accomplishing the goals of the project.
Step 4: Act. Conduct the experiment and analyze results
to know what you should do next
Training Packet
2016
Name: _____________________
Objectives:
-Become familiar with equipment and
reagents in the lab
-Review how to make solutions
-Become familiar with PubMed
-Become familiar with planning, conducting,
and recording experiments.
-Purify, quantify, and digest plasmid from
bacterial cultures.
-Induce and document production of
recombinant protein in E. coli.
-Learn the importance of experimental
controls.
-Become familiar with Good Laboratory
Practices.
Please read through entirely:
Training
- Lab Notebooks
- Pipetting
- Autoclave
- PubMed
- Making solutions
Evaluations and “Grading”:
- You will be evaluated on your performance in the lab as an employee of InnovaBio.
-
We ask you to report an honest assessment of yourself using the guidelines found in
the syllabus. You will be sent a link through Google Groups. Please complete these
in a timely fashion. The grading criteria can be found on page 14. Please review it.
You will need to fill out an evaluation upon completion of the training packet if not
sooner.
All significant dates and deadlines for evaluations, presentations, and assignments
will be noted on the Google Calendar.
PubMed and PubMed Central
PubMed is an online site set up and run by the government and the U.S. National Library of
Medicine. It contains sources from MEDLINE, science journals, and biomedical articles that
date back to the 1950’s. The site contains information on genomes, proteins, nucleotides, and
much more. The site is useful in that it provides researchers with up to date information about
genes, proteins, and other research projects going on. Some of the journals and articles within
PubMed are free, but many are abstracts of certain articles that can be purchased to view.
PubMed can be accessed by going to www.pubmed.com.
PubMed Central is an archive of life science journals, biomedical articles, and other
informational literatures that can be found on PubMed, but are full text and free to the public.
PubMed Central can be accessed through the PubMed website or by going to
www.pubmedcentral.com.
Your assignment is to research a topic of your choice on any of these two websites and print
out a primary research paper that seems appealing to you. Read through the article and write
a 1 page paper on the paper you have read. First, summarize the paper using the scientific
format (without using personal pronouns). Then, include what you learned, did not know, and
want to learn more about. This paper is due on the day you complete the training packet.
Proper Labeling of Sample Tubes
Proper labeling of sample tubes is a good lab habit that allows researchers to promptly
locate and identify the content of a sample tube. The label on a sample tube is used to guide a
researcher or a group of researchers involved on the same project to information regarding the
sample in question. This information typically consists of a detailed explanation about the origin
and nature of the sample. The exact tube label and specifications about the sample must be
written on the project notebook such that the researcher can correctly match the label on the tube
with the provided description. By following the labeling guidelines described below, a group of
researchers working on the same project can have access to samples at any given time along the
project workflow regardless of who obtained the sample.
What you did
Name on tube cap
What indicates
Name on side
of the tube
Labeling DNA
samples
You purified from 5
pHMGB1
different E. coli
1
colonies a pET21a(+)
4/4/10
plasmid with hmgb1
cloned in it (hmgb1 is
the gene of interest,
also called insert). You
did the plasmid prep on
April 4th, 2010.
Indicates hmgb1 cloned into
plasmid “p” was prepped on
April 4th, 2010 (you should
specify on your book the
identity of plasmid “p”).
On this date, your book
should have an entry
mentioning you prepped
plasmid pHMGB1 from
several different colonies.
Your initials.
The
concentration
of the
plasmid.
On April 4th, 2010, you
amplified then purified
a PCR product. The
gene you are working
with is hmgb1
HMGB1
pure PCR
4/4/10
Indicates hmgb1 from a PCR
experiment I conducted on
April 4th, 2010, is in solution
and that it is pure. On this
date, your book should have
an entry describing how you
amplified and purified hmgb1.
Your initials.
If you can
spare product,
then
determine and
write its
concentration.
On April 4th, 2010, you
digested your hmgb1
PCR product with
enzymes HindIII and
XbaI.
HMGB1
PCR
HindIII/XbaI
4/4/10
Indicates PCR product hmgb1
was digested with enzymes
HindIII and XbaI. Your book
should have an entry
describing how you digested
your PCR product on the
indicated date.
Your initials.
What you did
Name on tube cap
What indicates
Name on side
of the tube
Your initials.
On April 4th, 2010, you
purified your hmgb1
PCR product digest.
HMGB1
dig HindIII/XbaI
pure
4/4/10
Indicates you purified the
PCR product you digested
with HindIII and XbaI. On
the indicated date, your book
should have an entry
mentioning you purified your
hmgb1 PCR product digest.
You digested
pHMGB1-1 with
enzymes HindIII and
XbaI on April 4th,
2010.
pHMGB1-1
HindIII/XbaI
4/4/10
Indicates plasmid pHMGB1,
which comes from colony 1,
was digested with restriction
enzymes HindIII and XbaI on
April 4th, 2010. On this date,
your book should have an
entry describing how you
prepared the digestion(s).
On April 4th, 2010, you
ligated digested
plasmid and a digested
DNA fragment.
Name of plasmid
Name of DNA
fragment
lig
4/4/10
Could go like this:
pET21a(+)
hmgb1
lig
4/4/10
Indicates you ligated plasmid Your initials.
pET21a(+) with hmgb1 on the
mentioned date. On that date,
your book should have an
entry describing how you set
up the ligation reaction.
Name of protein
product
9/15/10
exp 1
“exp 1” indicates that the
gene expression was carried
out under condition “1”,
which should be explained in
detail in your lab notebook.
There can be as many
“numbers” as testing
conditions for gene
expression. You may have
several different tubes, each
with the protein product
manufactured at different
expression conditions.
Your initials.
Labeling protein
fractions
On Sep. 15th, 2010, you
prepared cells in SDS
loading dye to run a
SDS-PAGE for
analysis of gene
expression
Your initials.
The
absorbance
value of the
cell culture
before it was
spun down.
On Aug. 24th, 2010,
you lysed cells and
separated the soluble
from the insoluble
fraction by transferring
the soluble fraction to a
new tube.
Name of protein
product
8/24/10
Sol
con 1
Name of protein
product
8/24/10
Insol
con 1
“Sol” and “Insol” mean
soluble and insoluble,
respectively. “con 1”
indicates the lysis conditions
employed to obtain the
samples. Detailed
information describing the
lysis conditions should be
written in your lab notebook.
Your initials.
Information to
trace this back
to expression
conditions.
You treated sample
“insol, con 1” with
urea and/or other
detergents, and
separated the urea
soluble fraction from
the urea insoluble
fraction by transferring
the former to a new
tube. You did it on
Aug. 25th, 2010.
Name of protein
product
8/25/10
U-Sol
U-con 1
Name of protein
product
8/25/10
U-Insol
U-con 1
“U-sol” and “U-insol” mean
urea-treated soluble and
insoluble fraction,
respectively. “U-con 1”
indicates the conditions
employed under the urea
treatment. Detailed
information describing the
protein solubilization
treatment with urea should be
written in your lab notebook.
Your initials.
Information to
trace this back
to expression
conditions.
You initiated the
purification of your
protein by loading it on
a Ni2+ IMAC (HisTrap)
or any other of our
columns (HiTrap Q,
Sephacryl S-200 HR,
etc). You collected
fractions (1…etc.)
from your column as
proteins were being
eluted. This was done
on Aug. 25th, 2010.
Name of protein
product
Ni
8/25/10
1
“Ni” indicates that the sample
was eluted from a Ni2+ IMAC
(HisTrap) and “1” is the
fraction number. This
number should match the
AKTA FPLC elution pattern
for the run. This labeling
system can be used regardless
of the column from which the
protein is being eluted.
Your initials.
You continued the
purification of your
protein by loading it on
a Q HiTrap. You
collected fractions
(1…etc.) from your
column as proteins
were being eluted.
This was done on Aug.
26th, 2010.
Name of protein
product
Q
8/25/10
1
”Q” indicates that the sample
was eluted from a Q HiTrap
and “1” is the fraction
number. This number should
match the AKTA FPLC
elution pattern for the run.
This labeling system can be
used regardless of the column
from which the protein is
being eluted.
Your initials.
Exercises
Fill in the blocks
What you did
Name on tube cap
pET21-a
VG1-1
Nde1/HindIII
On Sep. 12th, 2010, you
set up a PCR
experiment and
obtained a PCR product
using primers 1 and 2
and condition 3. The
name of the PCR
product is A LOT.
What indicates
Name on side of the
tube
Lab Notebooks
You will write all procedures into your lab notebook provided by InnovaBio. You must write out
the experiment before you start handling any reagents. Every experiment should be
recorded whether “it worked” or “didn’t work”. All lab notebook entries need to follow the
following guidelines:












Name - List names of all who participated.
Title - Title the lab procedure you are doing.
Date - Write the date or dates of the experiment
Project and book numbers- Indicated in provided space. (Training Packet=Project No.1)
Purpose - Write out why you are doing the experiment
Materials - List all the materials you use with lot numbers for reagents that have them.
Protocol - List all the procedures you will go through to complete the experiment. Give
specific details and show any calculations.
Observations - Write out everything that you observed throughout the experiment. List
any modifications made in the methods you listed, all unexpected results, and anything
that would be useful to someone following the same experiment. Also place any graphs,
tables, or printouts.
Any corrections must be indicated by crossing out the incorrect part with a single line,
writing the correct information with your initials and the date.
Conclusion - From your observations, conclude whether the experiment worked or not.
Explain why the experiment did or did not work and back your conclusion up referencing
your methods, observations, or other materials.
References - Make sure to write in any references used in your protocol. Did you use
any papers, SOP’s, or other materials? Reference all sources that you used so that
someone else can find the information.
Each day you and a witness will sign and date the pages of the lab book.
Good Laboratory Practices
Good laboratory practices (GLP) were developed as a set of quality standards to ensure the
validity of and confidence in experimental results derived from non-clinical research. The major
components of GLP center on Quality Assurance (QA) and include Standard Operating
Procedures (SOPs), statistical procedures for data evaluation, instrument validation, reagent
and materials certification, specimen and sample tracking, and documentation and
maintenance of records. While this is only a partial list, it is evident that applying these
standards to most lab work and research could prove quite beneficial. Use of SOPs will ensure
that a variety of procedures are done consistently and reliably. Ensuring that the equipment
you’re using is good working order -and properly calibrated if necessary- and leaving that
equipment clean and in good working order for the next user will benefit the lab as a whole and
bolster confidence in results generated with that equipment. Careful monitoring and
documentation of stock solutions will promote confidence in their use and allow problems to be
identified readily should they arise. When setting up experiments, be sure all necessary
controls exist such that your experiments can be appropriately and confidently interpreted.
Careful note taking and documentation of experimental procedures and results (your lab
notebooks) will ensure confidence in and reproducibility of those results. In the lab, certain
stock solutions will have log sheets associated with them such that appropriate SOPs are
utilized and the date they were made and the person who made them are apparent. You will all
be making these solutions at some point, so adhering to these simple guidelines should allow
you to have confidence in solutions made by yourself as well as others and additionally ensure
consistency in their use. Some equipment in the lab may have logs as well but will at least
have guidelines for use which will include shut down procedures such that the equipment is
clean and good working order for the next user. Importantly, take time to read over the lab
notebooks section of the training packet; the guidelines outlined will ensure that your
experiments are appropriately documented. GLP standards exist in many of the work
environments you’ll potentially enter and becoming familiar with them now will only facilitate
the ease with which you assimilate them later on. Additionally, some of the more basic
standards will definitely improve the quality of your work and allow you to have confidence in
the outcomes. Take a minute to look up GLP standards in order to familiarize yourselves
with some of the terminology and practices.
Risk Assessments
In addition to knowing and understanding the laboratory rules and expectations as outlined in
the safety orientation, it is extremely important to be aware of any hazards associated with the
specific experiments you are doing. Hazards can be identified by completing a risk
assessment form, like the example provided below.
Assignment
Choose an experiment from the training packet and complete a risk assessment form. Forms
can be found in the lab or on the lab computer. Chemical safety information can be found in
the material safety data sheets (MSDS) in the lab or online. Please consider the following
questions when completing the risk assessment:
Description of task/experiment
- What experiment are you doing, and does it require special conditions (heat, gas
etc.)?
Hazard identification: equipment
- What equipment will you be using? Are there hazards associated with it?
Hazard identification: chemical
- What reagents are you using, and what are their chemical hazards?
- What is the National Fire Protection (NFPA) fire diamond? What do the
colors/numbers mean?
- What role does each reagent play in the experiment?
Safety controls; precautions; waste disposal
- What protective equipment will you need?
- How will you dispose of any waste?
Name
Date
Location
Phone
Description of task/experiment
Repetitive task Services used: Water
Power
Gas Other Temp 100 oC Pressure…
Preparation and use of agarose gel for DNA purification.
Melt agarose in TAE buffer in a conical flask by microwaving. Once the gel is cool, add
ethidium bromide and pour into gel casts. Once the gel has set, load DNA samples, run the gel
and visualize the DNA using the gel dock.
Hazard Identification: Equipment
Microwave – burn hazard
Electrophoresis Power Pak power supply – high voltage.
Gel Doc – UV light
Fire
Reactivit
y
Other
Use/purpose of reagent/notes
Health
Hazard Identification: Chemical
Name
50X TAE
(Tris-acetate-EDTA)
0
0
0
agarose
0
0
0
Gel preparation and running buffer. Tris-acetate
provides ions to conduct the electrical current to
aid DNA mobility, and maintains the pH. EDTA
chelates divalent metal ions to inhibit the action of
DNase on the DNA samples.
Agarose is a polymeric polysaccharide that forms
pores upon setting in the gel form. Agarose density
determines the pore size. These pores act like a
sieve and allow the DNA to be separated by size.
Ethidium bromide
3
0
0
Ethidium bromide is a fluorescent dye used to
visualize DNA in agarose gels by intercalating
between the base pairs of the two DNA strands.
Safety Controls; Precautions; Waste Disposal
Use personal protective equipment (PPE): lab coat, gloves, and safety glasses/goggles.
Use caution when microwaving glassware as the glass and contents will be hot. Use heatresistant gloves to carry glassware.
Be aware of general electrical shock hazards when using the PowerPak power supply.
When using UV light source, protect all skin from UV exposure by using gloves, lab coat etc.
Protect eyes by using appropriate face shield.
Ethidium bromide is a mutagen: use PPE when handling. Use designated chemical waste; do not
dispose down the drain.
Lab Benches
You will be provided a lab bench. You are responsible for keeping your bench clean and for
ensuring that all equipment is in proper working order. You will test your pipettes for proper
function using the Artell. Remove, from your bench, any unlabeled bottles or old reagents.
Clean your bench and overhead shelves of any dust or residue. Acquire new autoclaved water
and LB broth from the cabinet. Label them with your initials and the date.
Every day when you arrive and before you leave, sanitize your bench by clearing it of any
equipment, emptying biohazard buckets, and wiping the bench top with 70% ethanol. This will
help to prevent contamination coming from those who may have used your bench while you
were away.
Experimental Section
Be sure to write in your assigned lab notebook. You will be evaluated on the criteria in
the Lab Notebook section above. You must include lot numbers of reagents,
mathematical calculations and record all data (concentrations, gel pictures (trimmed
with lanes labeled), OD600 readings, etc.). All of the experiments in this section rely upon
previous steps in the overall process. You will be working with an expression plasmid pET32a
and performing a series of quality control experiments to ensure that the plasmid is correct
before expressing the protein encoded by the plasmid.
Your Assignment: construct a flow chart of the overall process indicating the order of
experiments and how they fit together.
Plasmid Preparation
- Transform DH5 E. coli with pET 32a plasmid using heat shock method.
o Use sterile technique to avoid contamination.
o Find protocol in SOP book.
o Plate 100 L on LB plate containing appropriate antibiotic.
o Grow (incubate) overnight at 37oC in plate incubator.
- Prepare three 5mL LB cultures (containing the appropriate antibiotic) using a
single colony from your transformation for each tube.
o Initial antibiotic concentrations are 50mg/mL Amp or 25mg/mL Kan.
o Final concentration in the LB broth are 100g/mL Amp or 25g/mL Kan
o Use sterile technique to avoid contamination.
o Use transformed DH5 E. coli containing pET 32a plasmid
o Grow culture overnight at 37oC in shaking incubator.
- Purify pET32a plasmid using the Qiagen Spin Prep kit and included protocol. Only
purify plasmid from one 5 mL culture. Store the remaining two sample pellets in the
freezer. Write into your lab notebook the version of the Qiagen protocol used (see the
cover of the book) and the entire protocol.
- Use Nanodrop spectrophotometer to determine plasmid concentration.
DNA and Amino Acid sequences in the expression region of the plasmid:
Plasmid Restriction Digest: Restriction digests can be used to clone a fragment of DNA such
as the one you are designing in the above assignment. Another purpose is to determine that
you are working with the correct plasmid. Please confirm that you have purified pET32a by
choosing two restriction sites with cleavage products that can easily be identified on a 1%
agarose gel.
- Using your pET32a miniprep, perform two single restriction digests with each enzyme
as well as the double digest with both enzymes. Finally include a no enzyme negative
control reaction. Use the table below to organize the reaction set-up. Rewrite the table
in your lab notebook.
o Use 250-500 ng pET32a plasmid for each digest.
o Use plasmid map (above) to determine appropriate enzymes and expected sizes
of bands.
o Incubate for 2 hours to overnight in 37oC water bath.
Reagents
pET 32a
ddH2O
10 X Buffer (#)
Enzyme 1
Enzyme 2
Total volume
Enzyme 1
Enzyme 2
Enzymes 1& 2
Neg. Control
Preparation of agarose gel materials:
- Prepare 1L of 1XTAE from the 50XTAE stock solution. Label and keep this stock at
your bench for your own use. You may want to test your 1XTAE by pouring a 1%
agarose gel and running 5µL of 2-Log DNA ladder.
- Refer to protocol in SOP notebook for making agarose gels. Use the appropriate
percentage of agarose and include 1µL of 10mg/mL Ethidium Bromide for 50mL
agarose. Caution: Ethidium Bromide is a mutagen. Always wear gloves when
handling this reagent and all the gel running equipment.
- Plan out your lane organization and write it in your lab notebook.
- Use 2-Log DNA ladder key to determine the product sizes.
- Include the resulting gel in your lab notebook. Make sure it is comprehensively
labeled and that you describe the results and conclusion in your lab notebook.
Assignment:
Often when working with a protocol given to you, you may encounter mistakes. You
should check protocols to ensure they will accomplish the intended result. The following is a
restriction digest to identify that the plasmid is correct. Find and correct the errors in the
following protocol. You will likely need to use on-line resources or resources in the lab.
Materials: 100ng/µL Plasmid pBlueScript KS+(pBS KS+), Restriction enzyme
NdeI, 10XNEB buffer 4, ddH2O
Procedure:
1. Calculation of the number of microliters needed for 350ng of plasmid.
350ng X ___µL
= 5µL
100ngpBS KS+
2. Restriction digest set-up. One reaction with no enzyme one reaction with
enzyme
ddH2O
pBS KS+
10XNEB buffer 3
NdeI
Total
30 µL 30 µL
5 µL 5 µL
3 µL 6 µL
0 µL 2 µL
38 µL 43 µL
3. Reaction conditions 25oC overnight.
4. 5% agarose gel Lane1
Lane2
Ladder
pBS KS+
No enzyme
Lane3
pBS KS+
With enzyme
PCR of the expression region using pET32a as template
- Dilute pET32a miniprep 1:5 in water for use as the template in a temperature
gradient PCR. Prepare a master mix for nine identical reactions which will vary in
annealing temperature. Only eight complete reactions are needed. The extra is to
have a little extra volume for small errors in pipetting accuracy.
- The following is a PCR recipe (add reagents in the listed order). Use the following
table to plan out the experiment. Determine the volumes required for a master mix
of nine reactions, and determine the final concentration of each reagent in the PCR
reaction. Rewrite the table in your lab notebook.
Reagents
1rxn (uL) 9rxn (uL) Final Concentration
Autoclaved dH2O
34.8
----------------------------10x Taq buffer
5.0
MgCl2 (25 mM)
5.0
Forward Primer (10 µM) (T7 promoter)
1.5
Reverse Primer (10 µM) (T7
1.5
terminator)
dNTP (10 mM each)
1.0
Diluted pET 32a
1.0
Taq Polymerase*
0.2
----------------------------Total
50
_______
Aliquot the master-mix into 8 separate PCR reaction tubes.
- Thermal Cycling Protocol (times are in minutes:seconds) You will need to use the
BIORAD thermal cycler with the single 96-well block.
1 cycle of: 94oC for 1:00
30 cycles of: 94oC for 0:30, 52-68oC (gradient) for 0:30, and 72oC for 1:00
1 cycle of: 72oC for 5:00, and 4oC final hold
- It is best to place PCR tubes in the center of the thermal cycler for most accurate
temperatures. For your reactions to match the gradient of temperatures indicated on
the thermal cycler place the tubes in a row of wells from back to front (rows A-H).
- Use the plasmid map to determine the expected size of the PCR product.
- Determine from the 1% agarose gel, the best annealing temperature for the T7
primer set.
Two assignments:
pET 32a is a circular plasmid from E. coli that contains the pET expression system. Lac operon
components are used to express a protein of interest when properly cloned into this vector.
The protein can be tagged with peptide/protein sequence that can aid in purification and in
making the protein soluble. You will need to complete two assignments.
1. Use NCBI to find a human protein of your choosing to theoretically clone into the
pET32a plasmid. Be sure to find the mRNA or cDNA sequence rather than genomic
sequence. You will design primers to the open reading frame (ORF) for PCR
cloning your selected protein coding sequence. Include on each end, a restriction site,
from the multiple cloning site region of pET32a. Please ensure that your protein product
will be in frame with the N-terminal Trx-tag and contains a stop before the C terminal
His-tag.
2. Please investigate the source and purpose of all DNA sequences highlighted in the box
below with bold lettering. Write a brief description of each component and how each
is used in this protein expression system. Submit with your completed training
packet.
Protein Expression
Day 1:
1. Transform BL21 (DE3) E. coli with your pET32a plasmid using the heat shock
method. Again you will need the appropriate antibiotic for pET32a.
Day 2: (Days 2 and 3 must be completed in succession. Ask for assistance if needed.)
2. In a 250mL Erlenmeyer flask, inoculate 50 mL of LB/amp using one colony of the BL21 E.
coli which has been transformed with pET32a plasmid. Culture overnight at 30°C in a
shaking incubator. This is your Overnight Culture.
3. Pour two 15% SDS-polyacrylamide gels (protocol in SOP notebook). Please view
the video on the following link if you have never poured polyacrylamide gel.
http://www.youtube.com/watch?v=EDi_n_0NiF4
Day 3:
4. Inoculate 50 mLs of LB/amp with 1 mL of your overnight culture. This is your
Expression Culture. Culture about one hour at 37°C in a shaking incubator.
Ideally you want the OD600 measurement in step 6 to be between 0.4 and 1.0.
5. Save 1mL of overnight culture to make a 15% glycerol stock. Mix 1mL of culture
with 165L of sterile glycerol. Store at -80oC in “Training Packet Box”. Dispose of
the remaining overnight culture by mixing with bleach then pouring down the drain.
6. Measure and record the OD600 of two 1 ml samples of the expression culture. Use a
disposable plastic cuvette and remember to first blank spectrophotometer with LB.
Save the cells in the cuvettes for step 8.
7. Induce remainder of the expression culture by adding 50 µl of 0.5 M IPTG, and
culture 2-4 hours at 37°C in a shaking incubator, or overnight at room temperature
on shaking platform.
8. Remove 900µL of the culture from the cuvette in step 6, transfer to a microcentrifuge
tube, then spin for 1 min at 14,000 rcf. Discard supernatant and store one sample at
-20 oC. Re-suspend the remaining pellet in 80 µl of 3X SDS sample buffer and 10µL
of 1M DTT. Incubate the cells and sample buffer in a boiling water bath or 95oC heat
block for 5 min. Label tube with sample description (eg. “un-induced cells”), E.coli
strain, plasmid, date, and your initials. Store sample at -20°C for SDS PAGE.
9. After 2-4 hours of induction at 37oC (or room temperature induction overnight:
Day 4), measure and record the OD600 of the induced expression culture. You will
likely need to dilute the culture 1:10 before measuring the OD600 (an OD600
measurement over 1 is inaccurate). In the cuvette, mix 900µL of LB broth with
100µL of expression culture then measure the OD600.
10. Centrifuge two 900µL samples of the expression culture, from the flask, for 1 min at
14,000 rcf. Prepare samples as for the un-induced samples in step 8. Label this tube
“induced cells” along with other relevant information.
11. Calculate the amount of resuspended cells to load on 15% SDS PAGE. An OD600 of
0.9 is the ideal cell density to load 10µL on the SDS PAGE gel. To determine the
volume of your samples to load, use the following equation: 0.9/measured OD600 X
10 = volume of sample to load on the gel. If necessary, make sure you multiply
any OD600 measurement by the dilution factor.
12. Run an SDS-PAGE gel using the calculated sample volumes. Remember to use a
protein ladder, usually 5 ul of Invitrogen’s Benchmark protein ladder.
13. Stain and destain the gel using the Quick Stain method. See the top of the
microwave for the protocol. After the gel has been sufficiently destained, dry the gel
between cellophane sheets.
14. Determine the expected mass of the expressed protein from the pET32a cloning and
expression region. Remember to determine the number of codons in the coding
sequence. Please use the average mass of an amino acid (110Da) to estimate the
expected size.
15. Lastly, determine the size of the expressed protein as it runs on the gel. Make a plot
of the distances each protein in the protein ladder runs from the top of the separating
gel versus the mass of each protein. From the plotted data, determine the equation
for the logarithmic regression curve of the ladder standard. Use the equation to
calculate the mass of the expressed protein from the distance it travels in the gel.
Place the dried gel and the plot in the results and conclusions section respectively.
Training Packet Completion checklist:

Read a primary paper (with experimental data) and write a one page summary.

Tube labeling exercise

Experimental flow-chart

Risk assessment

Corrected restriction digest protocol

Primer design for cloning of a human gene of your choosing into pET32a such that the
N-terminal Trx-tag is present in the synthesized protein.

Define and explain the significance for the bold words in pET32a box.

Lab notebook with all of the experiments and recorded data. You must include results
and conclusions.
Lab Math
What you need to know in order to make
and dilute solutions in the lab on a daily basis.
Name: _____________________
Converting Units
Explanation
When converting from one form of measurement to another, it can be difficult to know whether you are
doing things correctly. By setting up equations in the form of fractions (numerator on top, with
denominator below), you can be sure that you are performing the right mathematical operations to
convert units. This technique can be helpful in the lab as well as outside the lab.
A conversion factor is used to convert from one unit of measurement to another, and can be derived by
simple algebraic manipulations of an equality like 1 foot = 12 inches. That equality can be converted to
a fraction by dividing both sides of the equation by “12 inches”, which would result in a new equation:
1 foot = 12 inches which of course can be rewritten as 1 foot = 1
12 inches
12 inches
12 inches
By the same reasoning, you can divide both sides of the equation instead by 1 foot and end up with the
fraction inverted:
12 inches = 1
1 foot
Either way, you have just generated a conversion factor, which you can use to convert inches to feet, or
feet to inches, simply by multiplying the current measurement by the appropriate conversion factor,
which really is the same as multiplying by 1!
Example
1. There are 5280 feet in one mile (1 mile = 5280 feet). How many feet are there in 26.2 miles (the
distance of a marathon)?
26.2 miles x 5280 feet = 138,336 feet
1 mile
2. How many milligrams are there in 3.47 grams (conversion factor based on 1 g = 1000 mg):
3.47 g x 1000 mg
1g
Practice:
How many …
…liters in 1.496 mL?
…inches in 1.5 miles?
…milliliters in 0.89 liters?
…milligrams in 0.41 grams?
…microliters in 2.3 milliliters
…micrograms in 0.442 milligrams?
…milliliters in 5779 microliters?
…milligrams in 1985 micrograms
Moles
Explanation
One mole of a substance is equal to its formula weight measured in grams. The
abbreviation for mole is “mol”, all lower case letters (not much of an abbreviation, but
that’s the way it is!).
Example
Lead (Pb) has a formula weight of 207.2, so one mole of lead weighs 207.2 grams. How
much does 2 moles of lead weigh, or more correctly stated, how much mass does 2 moles
of lead have?
2 mol X 207.2 g = 414.4 g
1 mol
Carbon dioxide (CO2) has a formula weight of 44.0 (C=12.0, O=16.0, 12+16+16=44).
How much mass does 0.750 moles of CO2 have?
0.750 mol X 44.0 g = XXX g
1 mol
Practice
Using a periodic table, determine the mass, in grams, of 1, 2, 0.1, or 0.45 moles of the
following:
1 mole
Sodium (Na)
Chloride (Cl)
NaCl
Carbon (C)
Hydrogen (H)
Oxygen (O)
Oxygen (O2)
Carbon Dioxide (CO2)
Glucose (C6H12O6)
2 moles
0.1 mole
0.45 mole
Molarity
Explanation
The molarity of a solution is the number of moles of a substance in a 1 liter volume of the
solution. Molarity describes the concentration of a particular solute dissolved in a solvent
(solute in a solvent makes a solution). Molarity is abbreviated “M” (the abbreviation is
always capitalized).
Example
A 2.0 molar solution of NaCl (or any other molecule) contains 2 moles of solute,
dissolved in enough solvent to make a total volume of 1 liter of solution. How many
moles of NaCl are there in 400 mL of solution?
Remember: 2 molar = 2 M = 2 mol/L.
400 mL X
1L
X 2.0 mol = 0.80 mol NaCl (in 400 mL solution)
1000 mL
1L
Practice
Determine how many moles of solute there are in the following volumes of solution at
the stated concentrations:
How many moles of NaCl?
Molarity
1L
Volume (L)
0.2 L
3.5 L
1M
2M
0.1 M
0.45 M
Using the number of moles present in the above solutions, determine how many grams of
solute there are (FW of NaCl = 58.5).
Volume (L)
How many grams of NaCl?
1L
0.2 L
3.5 L
1M
2M
Molarity
0.1 M
0.45 M
Concentrations expressed in % (i.e. grams per 100 mL)
Explanation
Concentration is often described in terms of molarity, but can also be given in the grams
per 100 mL of solution, or percent. When the concentration of solute in solutions is
described as % weight/volume, or % w/v, that indicates that there are expressed number
of grams of solute in 100 mL of solution.
Examples
A solution of 1% (w/v) agarose consists of 1 g of agarose in each 100 mL of solution.
One hundred mL of 20% (w/v) SDS contains 20 g of SDS.
Agarose: 100 mL X 1 gram = 1 gram agarose
100 mL
SDS: 100 mL X 20 gram = 20 gram SDS
100 mL
Practice
Determine how much agarose is needed to make the following solutions:
% w/v
50 mL
150 mL
0.75 L
2L
0.8%
1.0%
1.2%
1.5%
Determine how much SDS is needed to make the following solutions:
% w/v
0.5%
1.0%
10%
20%
100 mL
200 mL
0.5 L
1.5 L
Making solutions of a particular molarity using dry
reagents
Explanation
Making a solution of a specific concentration is a fundamental skill any biotech
researcher needs to be able to do reliably. The concentration can be either weight/volume
(eg. grams/liter), molarity (eg. moles/liter), or even %w/v (eg grams/100 mL).
In order to calculate how much solute is needed to make a solution of a particular
molarity, there are three pieces of information that are needed: the desired volume and
concentration of solution, and the formula weight (grams/mole) of the reagent to be
dissolved in the diluent. You will have separate calculations for each substance.
Example
How much NaCl is needed to make 0.5 L of 0.9 molar NaCl?
0.5 L X 0.9 moles X 58.5 g = 26.325 grams
1L
1 mole
Practice
Prepare 300 mL of TE buffer, which consists of 10.0 mM Tris and 1.00 mM EDTA. The
formula weight of Tris is 121.1 and EDTA is 404.6.
Prepare 650 mL of TBS (Tris-buffered Saline), which consists of 100mM Tris, and 0.9%
NaCl
Prepare 425 mL of PBS, which consists of following reagents: 1.37M NaCl, 27mM KCl,
and 43mM Na2HPO4
Dilutions to a Specific Molarity
Explanation
Stock solutions of commonly used reagents are often used as a quick source of reagents,
but they usually need to be diluted to the working concentration (as the stock solutions
are generally at a higher concentration than the desired working solution). In order to
dilute those stock solutions, it is necessary to determine how much of the stock solution is
needed to make a solution containing the desired concentration of reagent (s). An
algebraic equation, C1V1=C2V2 is used to represent the Concentration (“C”) and Volume
(“V”). The subscripts are usually interpreted as “1” being the stock or more concentrated
solution, whereas “2” usually refers to the solution to be made, the working concentration
of solution.
If you know the concentration of the stock solution that you have available, and you
know the concentration and the volume of the solution you want to make, plugging
values into the algebra equation, and solving for the unknown variable, “V1”, allows you
to determine how much of the stock solution is needed to make the working solution.
Example
How much 20% SDS do you need in order to make 500 mL of 1% SDS?
C1 V1 = C2 V2
(20%)V1 = (1%) (500 milliliters)
V1 = 25 milliliters
Practice
How much 20% SDS would you need to make 50mL of 1% SDS, and how much water
would you need to dilute it with?
How much 2M NaCl do you need to make 0.5 L of 0.10 M NaCl, and how much water
would you need to dilute it with?
Dilution Ratios
Explanation
Some solutions in the lab are used with such frequency and volume, that it is easier to
store a reasonable volume of highly concentrated stock solution which can be easily
diluted. This is more practical than storing large volumes of working solutions, or making
up small batches of working solutions from powder each time they are needed, and the
stock solutions are often 4, 10, 50, or 100 times (4x, 10x, 50x, 100x, respectively) more
concentrated than the working solution.
In order to calculate how much of a stock solution is needed to make more working
solution, you first need to determine how much working solution you want to make. Then
simply divide the desired volume by the fold or “x” concentration of the stock solution.
Finally, bring the solution to the desired concentration using the appropriate diluent
(usually water).
Example
Make 1 L of 1X TGS (Tris/Glycine/SDS Running Buffer), using 5x TGS stock.
1000 mL (final volume) / 5 (concentration of stock) = 200 mL of 5X TGS needed, and
800 mL of water.
Practice
Make 200mL of 1X TGS from 5X Stock
Make 150mL of PBS from 10X Stock
Make 2L of PBS from 10X Stock
Make 1L of 1X TAE from 50X stock
Serial Dilutions
Explanation
Serial dilutions are used to reliably make multiple dilutions of a solution or sample, and
can be used to make extremely dilute solutions, far beyond the ability of any pipette to
dilute.
The process involves diluting the solution once, then using that dilution to make another
dilution, and repeating until the desired dilution(s) is (are) reached. The most commonly
used serial dilution schemes usually use 2-, 5-, or 10-fold serial dilutions.
The easiest way to set up serial dilutions is to pick an initial volume, and divide that by
the fold dilution you want. This is the most practical way, because the math is easy to
remember, you just have to make sure that when you do subsequent dilutions, there is
enough diluted sample left to use in your experiment!
Example
How would you make three 5-fold serial dilutions of BSA into 1mL of water?
First dilution (5-fold): 1mL (desired volume) / 5 (dilution ratio) = 200uL of BSA, to be
mixed with 800ul of water.
Second dilution (25-fold): 200µL of first dilution into 800µL of water. Since the first
dilution had 1000µL, and we just used 200µL of it, there is only 800uL of the first
dilution left!
Third dilution (125-fold): 200µL of second dilution into 800µL of water. Now we have
800µL of first and second dilutions, but 1000µL of the third dilution. This is not a
problem, unless you needed more than 800µL of the first two dilutions!
Practice
Make three 4-fold serial dilutions with an initial diluted volume of 400µL.
Make four 10-fold serial dilutions with an initial diluted volume of 1000µL
Dilutions Ratios
Explanation
Dilution ratios may be expressed as a combination of parts of solution mixed instead of
diluents into total volume. This alternate method is less common and is not standard
practice, but you should be aware of it. Some reagents used in the lab use this method for
making dilutions.
Step 1: Determine the amount of diluted sample you want to end up with.
Step 2: Next, determine the dilution ratio needed, and determine the ratio of
sample:diluent this will require. For 1:1 dilutions, you would mix sample with an equal
amounts of sample and diluent. For 1:4 dilutions, you would mix 1 part of sample with 4
parts diluent. For 1:9, you would mix 1 part of sample with 9 parts diluent.
Step 3: Determine the amount of sample and diluent to be used by multiplying the
desired volume of diluted sample (determined in Step 1) by the sum of the two numbers
in the ratio. 1:1 yields 2, 1:4 yields 5, and 1:9 yields 10.
Step 4: Divide the total volume needed by the sum of the ratio numbers. This value is
the amount of sample. Then multiply the amount of sample by the value representing the
portion of the diluents. For 100mL of a 1:4 part ratio: 100mL/5= 20mL of sample,
20mLx4= 80mL diluent
Doing it this way, the amount of diluted sample that is left is exactly what you wanted it
to be.
Example
Step 1: The amount of diluted sample needed is 500mL.
Step 2: Make a dilution of your sample that is 1:3 parts sample into water.
Step 3: 1 part + 3 parts = 4 parts
Step 4: Divide 500mL/4 = 125 mL. Therefore, 125mL sample and 125mLx3 = 375mL
diluents.
Practice
Prepare 500mL of a1:9 ratio of bleach to water. Show how much bleach and water you
will mix.
You will need to apply 3mL of western blot reagents. The instructions say to mix
solutions A and B in a 1:1 ratio. How much of each solution will you mix?
Dilution ratios (continued)
Explanation
Preparation of samples in the lab may also require you to dilute your concentrated stock
solution into your sample. This is especially true for preparing DNA samples to run on
an agarose gel. One part of the diluted sample comes from the stock solution while the
rest of the diluent comes from the sample. This calculation is a modification of the
C1V1=C2V2. In this case, the volume of V2 is the sum of the sample and the added
concentrated stock solution. V2= (VS + V1). Therefore C1V1=C2(VS+V1)
Step 1: Determine the amount of sample you have VS (e.g. 36µL).
Step 2: Determine the concentrated stock solution C1 (e.g. 3X) and the final
concentration C2 (e.g. 1X).
Step 3: Solve for V1. This number is the amount of concentrated stock you will need to
add to your sample.
Example:
Determine the amount of 5X buffer to add to 36µL of sample.
1. 36µL of sample.
2. 5X V1= 1X (36 µL +V1)
3.
5XV1=1X36 µL + 1XV1
5XV1-1XV1=1X36 + 1XV1- 1XV1
4XV1=1X36
V1=1X36/4X
V1= 9 µL
9 µL of 5X added to 36 µL of sample.
Practice:
1. Prepare 50µL agarose gel samples using 6X Loading buffer. How much loading
buffer will you need?
2. Prepare a glycerol stock of 2mL bacterial culture by mixing in10X concentrated
glycerol. How much 10X glycerol will you add?