An off-the-shelf, authentic and versatile undergraduate
molecular biology practical course.
David E. Whitworth
Institute of Biological, Environmental and Rural Sciences, Aberystwyth
University, Ceredigion, SY23 3DD, UK.
Email: [email protected]
Phone: +44 (0)1970 621828
Running Title: An off-the-shelf molecular genetics practical course.
Keywords: Education, Practical course, Molecular biology, Bacterial genetics
Abstract: We provide a pre-packaged molecular biology course, which has a broad context and is
scalable to large numbers of students. It is provided complete with technical setup guidance, a
reliable assessment regime and can be readily implemented without any development necessary.
Framed as a forensic examination of blue/white cloning plasmids, the course is a versatile
workbench, adaptable to different degree subjects, and can be easily modified to undertake novel
research as part of its teaching activities. Course activities include DNA extraction, RFLP, PCR, DNA
sequencing, gel electrophoresis, and transformation, alongside a range of basic microbiology
techniques. Students particularly appreciated the relevance of the practical to professional practice
and the authenticity of the experimental work.
Introduction:
In science education, a commonplace mode of teaching is the practical class, which usually manifests
as a laboratory-based experiment. The development of new practical classes is often timeconsuming, costly and reliant on the experience of staff members. However of all teaching events,
the learning outcomes of practicals are typically the most closely aligned with the future professional
practice of students. Practical classes are also particularly amenable to sharing between institutions,
as development of a practical can be separated from its implementation, much as experimental
protocols can be developed in one laboratory, and implemented in another.
In the UK, a Bachelor’s degree in one of the sciences will traditionally have different teaching
objectives in each academic year. The first year provides a broad background of principles and
knowledge, to ensure all students have a similar core knowledge entering year 2, regardless of their
academic background. The second year is focused on providing fundamental degree-specific
skills/knowledge, and students become relatively specialised in their final year. Laboratory-based
classes are usually employed as a teaching method in years 1 and 2, with experiments designed to
reinforce the theory acquired from lectures/books, and to provide practical experience. In the third
year, there are fewer taught practical classes; instead students pursue an extended novel research
project, culminating in production of a dissertation/thesis as a summative indicator of the ability to
undertake research.
Practical classes tend not to change year-on-year. This is often partly due to the large investment
required in their development (generation of experimental materials, training of technical support,
costs of equipment, materials and technical staff time, etc). Inertia also comes from the relatively
slow movement of the first and second year curriculum, meaning that the theory underpinning a
practical will not quickly be outdated, even if the experimental approach adopted does. The
experimental methods used in practical classes typically focus on a few versatile techniques, broadly
used in the discipline (including spectrophotometry, aseptic technique, enumeration of microbes,
pipetting, titration, gel electrophoresis, etc., in the case of microbiology and biochemistry). In
comparison, the techniques used in contemporary research are necessarily far more diverse,
sophisticated and rapidly advancing, but expense and time constraints preclude training students in
all those methodologies during their degrees.
As a consequence, the experiments undertaken in classes cannot be considered as novel research
and student classes are only rarely seen as partners in research (although examples are emerging,
for instance see [1-3]). The methods employed in teaching laboratories lag behind the methods used
in research laboratories. The relevance and ‘authenticity’ of laboratory classes is thus reduced, and
for many students, the 3rd year project can be their first experience of true research, and of many of
the techniques that they learn about during lectures.
Here we provide all the guidance and written materials needed to implement and assess a practical
course on molecular bacteriology, primarily aimed at undergraduate students. The course comprises
a week-long series of experiments, followed by a series of tutorials, and uses molecular biology
methods to investigate the differences between two bacterial strains. The methods employed are
used routinely in modern research laboratories, and the biological principles addressed have
relevance across many subject areas. Therefore the course can be easily inserted into most
biological sciences syllabuses, at multiple levels, and will give students an authentic experience of
molecular biology research.
The course is also designed to be versatile, allowing facile modification to align with staff
expertise/interests or with institutional requirements. It can be trivially amended in numerous ways
so that the students can even generate novel research data/materials which can supplement the
research activities of the academics involved.
Curriculum:
Course design. The course we are presenting (Molecular Lab Skills) was designed to provide practical
experience of modern molecular biology methods, but needed to be relevant to a wide range of
biological sciences degrees, including microbiology, biochemistry, genetics, zoology, aquatic biology
and plant science. Learning outcomes for the course included the acquisition of generic laboratory
skills, such as the following of instructions, accurate record keeping, and scientific presentation, as
these are particularly useful skills for future practice across the life sciences.
The course began with a week of daily laboratory sessions. During the week students undertook a
series of linked experimental procedures, as directed by a protocol booklet (supplemental material),
with each day’s practical producing material required for the next day’s experiment. This prevented
students forgetting what they had done during the previous practical session, which is important for
maximising engagement and attainment [4]. The experiments were framed as an investigation,
which gave the practical sessions continuity, and an atmosphere more akin to a research project
than a traditional practical class. Over subsequent weeks, the students (who each had an academic
tutor related to their degree scheme) met in groups with their tutors to discuss the methods
employed, the data obtained, and ways to interpret and present the experimental results. This
approach reflects that of a ‘flipped classroom’ as described by McLaughlin et al. [5], wherein
students are responsible for self-learning, with staff-contact promoting active learning methods.
Tutors were aided by the provision of a brief overview of the theory behind the investigation
(supplemental material).
Students’ attainment on the course was assessed by the production of an individually-written report,
and a piece of work produced collectively by the tutorial group. Tutors set the parameters of the
assessments for their tutees, manifesting an ‘assessment for learning’ approach [6], and were
encouraged to be innovative with the medium of the group work. Thus tutorial groups variously
created music videos, posters, documentaries, pieces of art, molecular models, even poetry, but
always related to the experimental work they had undertaken. Indulging the creativity of students is
important as objects of our own creation are more highly valued than those created by others [7].
Therefore the more creative the assessment, the more students will invest in the activity, promoting
their engagement, motivation and achievement.
The course splits the ability to observe and record data (as developed in the laboratory practical)
from the ability to interpret and report data. Model data is provided (Figure 2), so that students can
compare their data with expected results, and can meaningfully engage with the assessment even if
their experiments fail. Such provision of objective feedback is important in creating expertise and
preventing overconfidence [8, 9].
Practical sessions. The experiments used molecular biology methods to investigate two plasmids
(pBlue and pWhite) propagated in Escherichia coli. The two plasmids were initially generated using a
standard PCR product cloning kit using plasmid pCR2.1 [10]. PCR products were ligated into a
linearised plasmid by a conjugated topoisomerase, making use of complementary T/A overhangs of
the PCR product and plasmid [11, 12]. This single-pot reaction was then used to transform
chemically competent E. coli (Top10), with subsequent plating onto selective growth medium
containing X-gal. After overnight incubation, a mixture of white and blue colonies were obtained.
Colonies containing pBlue are coloured as the plasmid has recircularised without ligating a PCR
product insert, and linearisation recreates a functional lacZ gene. The LacZ product complements
the host’s chromosomally encoded mutant LacZ, producing a functional enzyme which hydrolyses Xgal into a blue pigment [13]. Conversely pWhite plasmids contain an inserted PCR product, which
disrupts the lacZ gene and host cells remain colourless (Figure 1). The experiments undertaken by
the students are aimed at providing a characterisation of pBlue and pWhite.
In our current version of the module, we divide the experiments over four days with a mid-week gap
(a detailed protocol booklet is provided as supplemental material). On Day 1, students use aseptic
technique to inoculate small cultures of Top10 [pBlue] and Top10 [pWhite] for overnight incubation,
and they also prepare selective nutrient agar plates containing X-gal. On the second day, plasmids
are extracted from the overnight cultures using ‘miniprep’ kits as provided by a variety of suppliers,
and used to set up three reactions. Purified plasmid DNA is digested by incubation with restriction
enzyme EcoRI, it is used to set up a reaction for nucleotide (ABI) sequencing, and also used to create
a PCR reaction mixture (using primers designed against plasmid sites flanking the PCR product
insertion site). PCR reactions and nucleotide sequencing are undertaken by module staff during the
mid-week gap day so that on Day 3 students return to the lab and are given their PCR reactions and
nucleotide sequence chromatogram files. The students then pour an agarose electrophoresis gel,
load their samples (restricted plasmid DNA, PCR reactions, and a DNA ladder) and subject the gel to
electrophoresis overnight. They also use their purified unrestricted plasmid DNA to transform
chemically competent Top10 and plate onto their selective X-gal plates. On the final day, DNA gels
are visualised by UV transillumination, and students count the number of blue and white colonies on
each of their plates.
Interpretation of data. Students finish the practical sessions having generated diverse types of data:
quantitative data (numbers of transformants), nucleotide sequence data, qualitative data
(phenotypes), and pictorial data (gel images). A typical gel image is shown in Figure 2.
From the gel image, students can infer that the two plasmids differ by the inclusion of an extra piece
of DNA, inserted between the two PCR primer binding sites, and they can estimate the size of
introduced DNA. Size estimation can be intuitive, or made more rigorous by plotting size of marker
bands against distance migrated, depending on the tutor’s whim. As well as providing a more
accurate size determination, plotting would also enable estimation of the plasmid backbone size,
which is deliberately larger than the largest marker band to allow discussion of issues around inter/extra-polation.
The colours of transformant colonies allow the students to infer that the plasmid DNA is responsible
for the colour phenotype, confirming the role of DNA as the genetic material. We collect together
every student’s results from the plasmid transformations and provide students with results for the
entire class. This allows a range of statistical tests and mathematical analyses to be performed.
Students can calculate means and measures of deviation, they can test for the significance of any
differences between the numbers of transformants for different plasmids, and they can identify and
test likely cases of mis-labelling. If transformations are performed with a dilution series, they can
test whether transformation frequency is dependent on the concentration of plasmid DNA, together
with measures of confidence.
Analysis of the nucleotide sequence of pWhite should confirm the students’ suspicions regarding the
nature and differences of the two plasmids. Sequences are provided as .ab1 chromatogram files,
which can be viewed using a downloadable program [14], and which allows tutors to discuss the
nature of dideoxy chain termination sequencing [15]. Students can extract the sequence data and
can check for the presence and distance between EcoRI sites in the plasmid, identify the insertion
site, and infer the size of the inserted DNA in pWhite. Many further analyses are possible using
public, free and intuitive web-services, depending on the tutor’s interests. For instance the inserted
sequence can be analysed using BLAST [16] to identify which organism the DNA came from, and
what gene the DNA encodes. Our version of pWhite contains an internal fragment from within the
gene encoding glyceraldehyde-3-phosphate dehydrogenase (GAPDH) of Myxococcus xanthus. The
behaviour of M. xanthus as a social developmental bacterial predator, the role of GAPDH in
metabolism and pathogenesis, and the crossover between these phenomena [17-19], provide lots of
starting points for interesting tangents for tutors to discuss with their tutees.
Outcomes. Because of the versatility and flexibility of this course, at Aberystwyth we concentrate on
the acquisition of general scientific skills by the students, including the abilities to follow an
experimental protocol, to record interpret and report experimental data appropriately, and to
present data/concepts through a variety of media. Students also gain an appreciation of the
importance of labelling properly, making accurate measurements and other practicalities of working
in a laboratory. This course was designed to maximise the acquisition of such skills, by providing
students with an intensive, week-long series of linked experimental procedures to undertake.
Concentrating on such generic skills keeps the course relevant to a broad range of degree subjects.
In addition, learners benefit from ‘belonging’ [20] and a prolonged period of collaborative work in
practical classes stimulates student engagement, motivation and appreciation of the subject area.
Importantly, the students also gain experience of a variety of subject-specific skills - working
aseptically, manipulating and cultivating bacteria, preparing growth media, and disposing of
contaminated material. They also learn how to extract DNA, manipulate DNA, analyse DNA and
introduce DNA into a bacterium. To complete the module successfully, students also need to acquire
conceptual knowledge and understanding of the nature of DNA, plasmids, restriction enzymes, PCR,
cloning, and blue/white selection. These techniques and principles have all stood the test of time,
being used in professional contexts for decades, with every probability of being used for decades to
come in most laboratories.
The course has now been implemented for four years (a total of 1001 students), as a first year
(second semester) module. The module gives reliable outcomes, with mark means and distributions
not changing significantly between years. The mean combined mark of the two assessments related
to the laboratory exercise is 62.5 %, ± 0.6 % with a standard deviation of 11.4 %, ± 0.2 %.
After the practical sessions we collected feedback on the experimental exercise by asking students
to rate statements according to a Likert scale (1 = strongly disagree, 5 = strongly agree). The highest
scoring statements across the cohorts were that ‘I could see relevance to my degree’ (mean score
4.7), and ‘I am enjoying the course’ (mean score 4.5).
Students were also given the opportunity to use their own words to praise one strength of the
course, and to criticise one thing they would change. The majority of responses were bland and
uninformative such as ‘it was fun’ or ‘interesting practicals’, however there were recurring themes
that hinted at educational benefits beyond mere entertainment. Perhaps unsurprisingly given the
prompting by the Likert statements, many respondents chose to praise the relevance to their
degree, but a further 9 % specifically made comments that could be paraphrased as ‘I could see the
relevance to professional practice’.
The lumping together of practicals into a week-long ‘project’ was selected for praise by 4 % of
respondants, and a further 4 % praised the ‘authenticity’ of the practicals, with comments such as ‘In
depth practical felt like real science’. Taken together, 7 % of students praised the practicals for being
‘challenging’, for using a ‘wide variety of techniques’, or for their ‘independence’ in undertaking the
practicals. Some individual praises were also encouraging as they showed our primary learning
outcomes were being achieved (‘taught good organisational skills’ and ‘showed importance in
following lab methods carefully’), and that the course was having a positive effect on the students’
perceptions of microbiology (‘I really enjoyed it. It's the first lab I've done with microbes’, ‘it was
useful as I was not thinking of going into lab work in the future’, ‘was good to see an application of
microbiology’).
Adopting the course. The practical elements of this course can be reproduced using commonly
available reagents, and a day-to-day list of the materials needed are provided as supplemental
information. Ideally, E. coli strains carrying plasmids pWhite and pBlue would be generated by
lecturers adopting this course, with bespoke inserts related to their own research. However the
strains described here are also available on request from the authors. The documentation required
to run the practical is also provided as supplementary material, including the protocol booklet, a crib
sheet for tutors and a troubleshooting guide.
Discussion:
Relevance: The effectiveness of learning experiences depends on the overlap with the future reality
in which the learning will be practised, and therefore learning benefits from being anchored in realworld problems [21,22]. Because of the fundamental nature of the techniques used here, it is trivial
to relate the experiments undertaken to aspects of modern living so that the students can see the
relevance of molecular biology.
The course introduces students generally to bacteria, and they can get an appreciation of the
appearance of bacteria, how quickly they grow, how pervasive they are, and how important aseptic
technique/contamination control is. Such insights can be extended straightforwardly to a
consideration of infectious organisms, and human efforts to reduce bacterial disease. In the
practical, selection for transformants uses antibiotics, which can be related to antibiotic therapy and
the increasingly publicised issue of antimicrobial resistance. Restriction enzyme digests with
visualisation by gel electrophoresis (essentially RFLP – restriction fragment length polymorphism)
also underpins the forensic fingerprinting techniques commonly seen in TV dramas,
paternity/maternity testing and neonatal testing for genetic disorders. One year, the running of this
course coincided with the UK horsemeat scandal, wherein horse flesh entered the human food chain
undeclared, and we were able to use that example to explain the importance of molecular methods
in identifying the taxonomic origin of DNA, and how it could be used to test for the provenance of
foodstuffs.
Authenticity: The course started with a week-long lab practical, which engendered the atmosphere
of a research project rather than an isolated practical. The experiments were linked together so that
it mattered how successfully the students implemented each day’s protocols. The experiments were
also framed as an investigation rather than as an abstract theory-driven experiment. This gave the
experiments a narrative and a real-world feel, showed the students how the methods could be used
in practice, and the use of molecular biology kits also contributed to the authentic feel of the course.
The use of story is one of Winne and Nesbit’s 25 heuristics for promoting learning [20] and helps
enhance the perceived overlap between learning and practice [22].
Students have been shown to value highly courses that they judge as authentic, and the ideal of
fairness in assessment is also strongly linked with perceived authenticity and relevance [23, 24].
Versatility: The laboratory sessions have been designed for use as a framework for further
exploration rather than as proof of established principle. Tutors are encouraged to steer tutorial
discussions onto topics of interest, even if only of tangential relevance. They are also able to tailor
the assessments as they wish, maybe to relate to their students’ degree specialisms, background or
career aspirations. For instance, for a more taxonomic slant tutors could use the DNA sequences to
undertake a phylogenetic analysis of organismal taxonomy/evolution.
This course is presented as a problem-based learning (PBL) exercise as challenging students’ initial
processing of information benefits their long-term learning [25, 26]. PBLs and other forms of
investigative, active-learning approaches (such as the flipped classroom approach) can be criticised
because they create a cognitive block by providing large amount of new information which is
incongruous with the students’ world view [27]. However, the real-life research process is full of
such cognitive blocks - experiments are often performed with only partial understanding of methods
or reagents, and novel data are often not as expected. Hindsight becomes vital in updating the
world-view and providing retrospective understanding, and such deep mental processing ultimately
stimulates greater learning [25]. In this course we wanted to engineer such a learning situation to
convey an authentic experience of the research experience, however an introductory lecture could
easily be provided to forearm students about the theory if desired.
Flexibility/Modifications: As the assessment is not inextricably linked with the practical sessions, it is
trivial to change the assessment regime without modifying the experiments. For instance adopters
may wish to develop an alternative assessment which focuses more on the acquisition of the
subject-specific skills.
The practical elements of this course can also be readily customised to allow the generation of novel
research data, or to fit with different disciplines. A simple change is to provide different versions of
pWhite to each pair of students/tutorial group, which then gives each pair ownership/responsibility
of their part of the practical. Different versions of pWhite can be generated with inserts that relate
to particular degree subjects. For instance, a photosynthesis gene could be cloned for plant
scientists, a developmental gene for zoologists, or a virulence gene for biomedical students.
A simple variation to the experimental scheme allows students to clone their own PCR products,
generating plasmids which could be of subsequent use in the staff members’ research. For instance
the pWhite we currently use was originally created as a gene disruption vector for use in
Myxococcus xanthus. In this variation we recommend setting up a PCR reaction and pouring plates
on day 1, cloning and transformation on day 2, inoculating cultures of blue/white transformants on
day 3, with DNA extraction and restriction on day 4. Day 5 would involve setting up sequencing
reactions and visualising PCR products and restriction digests by agarose electrophoresis. A microbial
ecology modification of this variant would be to use degenerate or universal primers to amplify
genes for cloning from metagenomic DNA, or from previously isolated environmental strains.
We have even run more extensive non-microbiological variations for students on different degree
schemes, including a ‘taxonomy’ variant primarily for zoologists and ecologists. In this variation the
students start by identifying a mollusc using a traditional identification key. They then use molecular
biology kits to extract the mollusc DNA, PCR amplify and sequence a conserved gene, and then use
phylogenetic approaches to confirm the identity of their mollusc. We found that tailoring variations
to particular degree schemes increased both student and staff engagement, and increased their
enjoyment of the practical.
Pedagogy: The development of this course was grounded in several important pedagogic principles.
In addition to meeting various subject-specific and general learning outcomes, it was very conscious
of the need to motivate students to engage with the experiments and assessments. This was
engineered through the deliberate focus on authenticity of experience, anchored in a real-world
problem. Ubiquitous experimental methods were linked together with a narrative thread that
immersed students in a challenging context requiring evaluation and synthesis of knowledge, the
highest levels of Bloom’s taxonomy of educational objectives [28]. Winne and Nesbit have proposed
25 heuristics for promoting learning, several of which were exemplified in this course, including the
generation effect, use of stories, feedback effects, deep questions and anchored learning [21].
Summary: Of the commonest types of teaching event, practical classes particularly lend themselves
to dissemination between institutions because of the ability to separate initial development from
implementation. Such dissemination allows institutions to offer practical courses to students which
are beyond the direct experience of their staff, saving staff time and enabling sharing of best
practice across the sector.
The course we are describing can be adopted easily, as we provide a full description of the course, all
course documents and technical notes. It provides an immersive student experience, of relevance to
their degree course, profession and modern life.
Supplemental Materials:
A variety of supplementary materials are available to allow straightforward adoption of the course.
The protocol booklet for students is provided, alongside a guide for tutors, a day-to-day description
of technical requirements including preparation guidelines, and a troubleshooting guide.
File pWhite.ab1 is a sequence chromatogram for plasmid White, and the annotated sequence of the
cloning region of pBlue is provided as pBlue.doc. Sample class data for the transformation
experiment is included (Class_data.xls), and a model gel image is provided as Figure 2 of this article.
Students will also benefit from direction to a free chromatogram viewer [11] and the cloning manual
for the kit that was used to create pWhite and pBlue [7].
Acknowledgements:
The assistance of Alun G.L. Evans is gratefully acknowledged for providing invaluable technical
support in the development and delivery of this course at Aberystwyth.
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Figure Captions:
Figure 1. Plasmids Blue and White. Insertion of a 0.8 Kbp PCR product into pBlue produced pWhite
which has a disrupted lacZ gene. The insertion increases the distance between the two EcoRI sites,
and also increases the size of PCR products obtained using the primers whose binding sites are
shown with arrows.
Figure 2. A typical gel image acquired by students. Lanes 1 and 2: EcoRI-digested pBlue and pWhite
respectively. Lanes 3 and 4: PCR reaction products generated using pBlue and pWhite respectively as
templates. Lane 5: DNA markers of known molecular weight (Hyperladder II, Bioline), allowing
estimation of the sizes of the DNA fragments.
Figures:
Figure 1.
Insertion site
EcoRI EcoRI
EcoRI
pBlue
4.0 Kb
pWhite
4.8 Kbp
AntibioticR gene
AntibioticR gene
Figure 2.
1
2
3
EcoRI
4
5
- 1000 bp
- 300 bp