Solution Manual For
Molecular Biology Techniques, A Classroom Laboratory Manual Edition 5 By Sue Carson,
Heather B. Miller, D. Scott Witherow and Melissa C. Srougi
Chapters 1-33
1 Discussion Question Answer Keys
2 Pre-Lab Questions Answer Key
Lab Session 1
1. What are some real-life applications of biotechnology? What are some important recombinant
proteins and/or recombinant organisms that are used today?
Student answers will vary.
2. What are your goals in taking this class? What are you hoping to learn, and how do you hope it
will expand your career or future research?
Student answers will vary.
Lab Session 2
1. If you do an absorbance reading after plasmid purification and get a A260/A280 of less than
1.8, how could you further purify the sample to get rid of the protein contamination? Is it
always necessary to have completely pure DNA? What are some cases where it would or
would not be?
There are a number of options to remove protein contamination. Phenol/chloroform extraction,
ethanol precipitation, chromatography steps (size exclusion or ion-exchange) are some of the
possibilities.
It is not always necessary to have pure DNA, depending on the goal of the experiment. For
example, unless there are protease present in the contaminating protein, extra protein will not be
a problem when performing a restriction digest. The buffer for restriction digest contains BSA (a
protein), so clearly the presence of protein is not a problem for this. However, if the DNA was
being used to transfect mammalian cells, the presence of proteins could severely hinder the
effectiveness of the transfection agent and limit the amount of DNA getting into the cells.
2. Why do you increase the pH to denature the plasmid and chromosomal DNA during
alkaline lysis rather than using high temperatures, which would also denature DNA?
, Because chromosomal DNA is much larger, the temperature would have to be quite high to
denature it. Heating a sample to such a high temperature and then cooling it back down to
reanneal, would take significantly longer than using a reagent that instantly changes the pH.
Lab Session 3
1. How efficient would PCR be if we set the annealing temperature higher or lower than the
calculated melting temperature? How would a higher or lower temperature affect the
annealing capability of the primers and the final quantity or quality of the products? How
would a gel of such PCR products look compared to PCR products obtained at the optimum
annealing temperature?
If the annealing temperature is lower than the calculated melting temperature, the binding of the
primers will be less stringent. In other words, binding of the primers is easier, but also more
likely to bind to undesired (non-specific) regions of the template. Overall, you would expect more
total product in this case. If the annealing temperature is higher than the calculated melting
temperature, the binding of the primers will be more stringent. This means there is likely to be
less product, but amplified products are more likely to result from specific priming.
Because there is likely to be more priming occurring at the lower annealing temperature, the
quantity is expected to be higher; however, the quality (in terms of specificity might be lower).
This would show up on an agarose gel as a more intense band of the specific band at the lower
temperature, but also perhaps the appearance of non-specific bands of other sizes. At a higher
temperature, you’re more likely to observe only one band at the correct size, although it might be
less intense.
2. People are most familiar with using Taq DNA polymerase, but in this lab, we are using a
different thermostable DNA polymerase. What is it and why are we using it instead?
Vent DNA polymerase is used because it has 3'→5' proofreading exonuclease activity that helps
to increase its fidelity.
3. Why can circular plasmid DNA appear as multiple bands on an agarose gel? Why doesn’t it
run at the same apparent size as linear DNA of the same length on the gel?
Circular plasmid can appear as multiple bands because supercoiled DNA and nicked (DNA with
a break in only one of the strands) migrate differently. Supercoiled DNA will migrate faster,
because the three-dimensional size is smaller. Nicked DNA runs much slower, as the circle has
more resistance going through the gel. Linear DNA will migrate somewhere between the nicked
and supercoiled DNA.
4. A different DNA ladder from NEB (Cat #N3200L) comes as 500 µL of a 1000 µg/mL
solution and is not pre-mixed with the 6× DNA loading buffer. If you want to load 150 ng of
, this ladder, how would you prepare the solution and what volume of the solution would you
load on your agarose gel?
There are many possible ways to do this that involve diluting the original sample. One such idea
would be dilute the original sample 1:100 (1 µL of the 1000 µg/mL + 99 µL of water). This would
result in a 10 µg/mL solution of the ladder. Then to load 150 ng, you would need to load 15 µL of
this sample. But first, it must be mixed with 6× DNA loading buffer. The correct ratio for a 6×
buffer is 5:1 (sample to buffer). So, mix 15 µL of the ladder with 3 µL of the 6× and load all 18
µL on the gel.
You would not want to mix 0.15 µL of ladder with 0.03 µL of 6× buffer because these volumes
are too small to pipette.
Lab Session 4
1. What are the potential reasons more than one band may appear when you run your PCR
product on a DNA gel?
Nonspecific binding of primers to template resulting in an undesired product.
Template could show up as a faint band, particularly if too much was used in the reaction.
Primers often show up as a smear at very low molecular weights at the bottom of the gel.
2. Would the cloning of the egfp gene into the pET-41a vector work if we skipped the
NcoI/NotI digestion of the PCR amplified egfp? How would the results of next week’s
ligation be affected?
No, it wouldn’t work. The NcoI/NotI digestion is imperative to create the sticky ends needed for
ligation. Without it, ligation would not occur and the two pieces (vector and insert) would remain
apart.
3. Normally, a few additional nucleotides need to be engineered 5′ of the restriction sites on
both the forward and reverse primers when cloning by PCR in the method we are using.
Why is this necessary? How can you determine how many additional nucleotides should be
added to the ends of the PCR primers? Does it matter which nucleotides are added?
The extra nucleotides are added to give the restriction enzymes something to bind to so they can
achieve their complete catalytic function. Not all enzymes need extra nucleotides on the end of the
sequence to efficiently cleave the DNA, but it never hurts to have them there. You can find out
which enzymes need how many nucleotides by using a table such as one provided by New
England Biolabs (Cleavage Close to the End of DNA Fragments; https://www.neb.com/en-
us/tools-and-resources/usage-guidelines/cleavage-close-to-the-end-of-dna-fragments). Finally, it
does not matter which nucleotides are added, since added nucleotides will not be part of the final
product.
, Lab Session 5
1. In our ligation, we used a 1:5 molar ratio of vector–to-insert. How could different
molar ratios (1:1 or 1:10) affect the ligase reaction? What are pros and cons of
excess vector versus excess insert?
The more insert present (ie, 1:10 ratio), the more total sticky ends are present to bind to
not only the vector, but also other inserts. You would be more likely to see concatemers of
multiple inserts in this case. With less insert present (ie, 1:1 ratio), there are less total
sticky ends, which would mean fewer concatemers, but also less ends to interact with the
vector to make the preferred product. Overall, the expectation would be that the reaction
would work regardless, with the number of vector plus a single insert changing slightly
across a wide range of ratios.
1. You perform a ligation with NcoI/NotI-digested, pET-41a vector and the egfp insert from
pEGFP-N1 (as in lab). You then transform the ligation mix into E. coli and plate on LB
medium containing kanamycin. Consider the following unexpected outcomes, and suggest
controls:
a. Nothing grows. What controls might you design to determine whether the E. coli
cells are viable (alive) versus whether the E. coli is competent? What would the
results of those controls be?
b. You see lots of growth, but when you isolate plasmid from numerous E. coli
colonies, all you find is pET-41a with no insert. Suggest the most likely way these
colonies arose (aside from contamination). What would the results of your controls
look like if your suspicion was correct?
a. If nothing grows, the most obvious control would be to make sure the cells are alive and
competent. To test for competency, transform cells with uncut pET41-a and confirm they
grow. To test for cell viability (confirming the cells are still alive, although they might not
be competent and able to take up foreign DNA, such as the plasmid), the cells could
simply be plated on LB media with no antibiotics. If the bacteria grow, they are viable. If
not, they are essentially dead cells.
b. The most likely result for a lot of colonies without insert is from pET-41a not being cut to
completion. Since supercoiled DNA transforms more efficiently, a small amount of uncut
pET41-a can lead to many colonies. If this were the case, it should be clear by looking at
the control where Ncoi/NotI digested pET41-a was transformed without insert or ligase.
This control should have no colonies, although often there are a few
Lab Session 6
1. Will the restriction mapping screen be able to detect transformants with multiple inserts?
Why?
Molecular Biology Techniques, A Classroom Laboratory Manual Edition 5 By Sue Carson,
Heather B. Miller, D. Scott Witherow and Melissa C. Srougi
Chapters 1-33
1 Discussion Question Answer Keys
2 Pre-Lab Questions Answer Key
Lab Session 1
1. What are some real-life applications of biotechnology? What are some important recombinant
proteins and/or recombinant organisms that are used today?
Student answers will vary.
2. What are your goals in taking this class? What are you hoping to learn, and how do you hope it
will expand your career or future research?
Student answers will vary.
Lab Session 2
1. If you do an absorbance reading after plasmid purification and get a A260/A280 of less than
1.8, how could you further purify the sample to get rid of the protein contamination? Is it
always necessary to have completely pure DNA? What are some cases where it would or
would not be?
There are a number of options to remove protein contamination. Phenol/chloroform extraction,
ethanol precipitation, chromatography steps (size exclusion or ion-exchange) are some of the
possibilities.
It is not always necessary to have pure DNA, depending on the goal of the experiment. For
example, unless there are protease present in the contaminating protein, extra protein will not be
a problem when performing a restriction digest. The buffer for restriction digest contains BSA (a
protein), so clearly the presence of protein is not a problem for this. However, if the DNA was
being used to transfect mammalian cells, the presence of proteins could severely hinder the
effectiveness of the transfection agent and limit the amount of DNA getting into the cells.
2. Why do you increase the pH to denature the plasmid and chromosomal DNA during
alkaline lysis rather than using high temperatures, which would also denature DNA?
, Because chromosomal DNA is much larger, the temperature would have to be quite high to
denature it. Heating a sample to such a high temperature and then cooling it back down to
reanneal, would take significantly longer than using a reagent that instantly changes the pH.
Lab Session 3
1. How efficient would PCR be if we set the annealing temperature higher or lower than the
calculated melting temperature? How would a higher or lower temperature affect the
annealing capability of the primers and the final quantity or quality of the products? How
would a gel of such PCR products look compared to PCR products obtained at the optimum
annealing temperature?
If the annealing temperature is lower than the calculated melting temperature, the binding of the
primers will be less stringent. In other words, binding of the primers is easier, but also more
likely to bind to undesired (non-specific) regions of the template. Overall, you would expect more
total product in this case. If the annealing temperature is higher than the calculated melting
temperature, the binding of the primers will be more stringent. This means there is likely to be
less product, but amplified products are more likely to result from specific priming.
Because there is likely to be more priming occurring at the lower annealing temperature, the
quantity is expected to be higher; however, the quality (in terms of specificity might be lower).
This would show up on an agarose gel as a more intense band of the specific band at the lower
temperature, but also perhaps the appearance of non-specific bands of other sizes. At a higher
temperature, you’re more likely to observe only one band at the correct size, although it might be
less intense.
2. People are most familiar with using Taq DNA polymerase, but in this lab, we are using a
different thermostable DNA polymerase. What is it and why are we using it instead?
Vent DNA polymerase is used because it has 3'→5' proofreading exonuclease activity that helps
to increase its fidelity.
3. Why can circular plasmid DNA appear as multiple bands on an agarose gel? Why doesn’t it
run at the same apparent size as linear DNA of the same length on the gel?
Circular plasmid can appear as multiple bands because supercoiled DNA and nicked (DNA with
a break in only one of the strands) migrate differently. Supercoiled DNA will migrate faster,
because the three-dimensional size is smaller. Nicked DNA runs much slower, as the circle has
more resistance going through the gel. Linear DNA will migrate somewhere between the nicked
and supercoiled DNA.
4. A different DNA ladder from NEB (Cat #N3200L) comes as 500 µL of a 1000 µg/mL
solution and is not pre-mixed with the 6× DNA loading buffer. If you want to load 150 ng of
, this ladder, how would you prepare the solution and what volume of the solution would you
load on your agarose gel?
There are many possible ways to do this that involve diluting the original sample. One such idea
would be dilute the original sample 1:100 (1 µL of the 1000 µg/mL + 99 µL of water). This would
result in a 10 µg/mL solution of the ladder. Then to load 150 ng, you would need to load 15 µL of
this sample. But first, it must be mixed with 6× DNA loading buffer. The correct ratio for a 6×
buffer is 5:1 (sample to buffer). So, mix 15 µL of the ladder with 3 µL of the 6× and load all 18
µL on the gel.
You would not want to mix 0.15 µL of ladder with 0.03 µL of 6× buffer because these volumes
are too small to pipette.
Lab Session 4
1. What are the potential reasons more than one band may appear when you run your PCR
product on a DNA gel?
Nonspecific binding of primers to template resulting in an undesired product.
Template could show up as a faint band, particularly if too much was used in the reaction.
Primers often show up as a smear at very low molecular weights at the bottom of the gel.
2. Would the cloning of the egfp gene into the pET-41a vector work if we skipped the
NcoI/NotI digestion of the PCR amplified egfp? How would the results of next week’s
ligation be affected?
No, it wouldn’t work. The NcoI/NotI digestion is imperative to create the sticky ends needed for
ligation. Without it, ligation would not occur and the two pieces (vector and insert) would remain
apart.
3. Normally, a few additional nucleotides need to be engineered 5′ of the restriction sites on
both the forward and reverse primers when cloning by PCR in the method we are using.
Why is this necessary? How can you determine how many additional nucleotides should be
added to the ends of the PCR primers? Does it matter which nucleotides are added?
The extra nucleotides are added to give the restriction enzymes something to bind to so they can
achieve their complete catalytic function. Not all enzymes need extra nucleotides on the end of the
sequence to efficiently cleave the DNA, but it never hurts to have them there. You can find out
which enzymes need how many nucleotides by using a table such as one provided by New
England Biolabs (Cleavage Close to the End of DNA Fragments; https://www.neb.com/en-
us/tools-and-resources/usage-guidelines/cleavage-close-to-the-end-of-dna-fragments). Finally, it
does not matter which nucleotides are added, since added nucleotides will not be part of the final
product.
, Lab Session 5
1. In our ligation, we used a 1:5 molar ratio of vector–to-insert. How could different
molar ratios (1:1 or 1:10) affect the ligase reaction? What are pros and cons of
excess vector versus excess insert?
The more insert present (ie, 1:10 ratio), the more total sticky ends are present to bind to
not only the vector, but also other inserts. You would be more likely to see concatemers of
multiple inserts in this case. With less insert present (ie, 1:1 ratio), there are less total
sticky ends, which would mean fewer concatemers, but also less ends to interact with the
vector to make the preferred product. Overall, the expectation would be that the reaction
would work regardless, with the number of vector plus a single insert changing slightly
across a wide range of ratios.
1. You perform a ligation with NcoI/NotI-digested, pET-41a vector and the egfp insert from
pEGFP-N1 (as in lab). You then transform the ligation mix into E. coli and plate on LB
medium containing kanamycin. Consider the following unexpected outcomes, and suggest
controls:
a. Nothing grows. What controls might you design to determine whether the E. coli
cells are viable (alive) versus whether the E. coli is competent? What would the
results of those controls be?
b. You see lots of growth, but when you isolate plasmid from numerous E. coli
colonies, all you find is pET-41a with no insert. Suggest the most likely way these
colonies arose (aside from contamination). What would the results of your controls
look like if your suspicion was correct?
a. If nothing grows, the most obvious control would be to make sure the cells are alive and
competent. To test for competency, transform cells with uncut pET41-a and confirm they
grow. To test for cell viability (confirming the cells are still alive, although they might not
be competent and able to take up foreign DNA, such as the plasmid), the cells could
simply be plated on LB media with no antibiotics. If the bacteria grow, they are viable. If
not, they are essentially dead cells.
b. The most likely result for a lot of colonies without insert is from pET-41a not being cut to
completion. Since supercoiled DNA transforms more efficiently, a small amount of uncut
pET41-a can lead to many colonies. If this were the case, it should be clear by looking at
the control where Ncoi/NotI digested pET41-a was transformed without insert or ligase.
This control should have no colonies, although often there are a few
Lab Session 6
1. Will the restriction mapping screen be able to detect transformants with multiple inserts?
Why?
