Information

Problem with joining the gene of interest into the open plasmid when their lengths are not the same


Before the recombinant plasmid is obtained in recombinant DNA, the gene of interest is inserted into a linearized plasmid by DNA ligase.

What happens if the length of the gene of interest is not the same as that of the gap in the open plasmid? (e.g. far longer) Can the gene still be connected with the gap or are other processes needed before the recombinant plasmid can be put back into the bacteria?


From your question I suspect that you are thinking of DNA as being a very rigid molecule - it isn't.

Double stranded DNA is reasonably flexible and a useful mental model would be to think of this as being like tying the ends of two pieces of thin wire together to make a circle.

Double stranded DNA can make a closed circles smaller than 250 bp long1,2 and I don't think there is any known upper limit - for example there is a known circular bacterial chromosome 14,782,125 bp in length3.

A typical plasmid can accommodate inserts of any size up to total size of around 50 kb, but plasmids that are more than 20 kb are very difficult to work with and may require special transformation techniques.

References:

1: Thibault, Thomas et al. “Production of DNA minicircles less than 250 base pairs through a novel concentrated DNA circularization assay enabling minicircle design with NF-κB inhibition activity.” Nucleic acids research vol. 45,5 (2017): e26. doi:10.1093/nar/gkw1034

2: Shore, David, Jorg Langowski, and Robert L. Baldwin. "DNA flexibility studied by covalent closure of short fragments into circles." Proceedings of the National Academy of Sciences 78.8 (1981): 4833-4837

3: Land, Miriam et al. “Insights from 20 years of bacterial genome sequencing.” Functional & integrative genomics vol. 15,2 (2015): 141-61. doi:10.1007/s10142-015-0433-4


Plasmid transformation is a mechanism by which we can introduce genes of interest into bacterial cells. In order to do this, we isolate another plasmid, insert the gene we want, and then put the plasmid back into the bacterial cells.

We can isolate the plasmid via various biochemical means, and plasmids are small enough such that we can isolate whole plasmids. (We have difficult isolating whole pieces of longer DNA, which tends to break up into fragments.) Once we have isolated the plasmid, we can apply restriction enzymes (such as EcoRI) for the specific sequences that we want to the plasmid and the gene of interest. The restriction enzymes cleave the plasmid and our gene at cut sites often, this leaves stick ends (ends of DNA with a single-stranded strand between two and four bases long). These sticky ends readily bond back together with other matching DNA pieces in this manner, we can join the gene of interest into our plasmid or cut other pieces out of the plasmid. After adding the restriction enzyme, we add DNA ligase to ligate the plasmid back together (to reform the covalent bonds that hold the DNA together).

Once we have our ligated plasmid, we would like to insert it into bacterial cells. Some bacteria are naturally competent, and will simply accept plasmids from their environment. However, many other bacteria (such as E. Coli), are not naturally competent, and we have to induce competence in order to get them to accept the plasmid. There are electrical methods to induce competence, which apply a field and essentially break the bacteria open in order to accept the plasmid although most of the bacteria die, some survive and form colonies. There are also chemical methods to induce competence, such as adding calcium chloride to E. Coli, but their mechanism of action is more complicated.

Finally, once we have inserted our plasmid into bacterial cells, we can grow them back into colonies. If we need to, we can apply selection for recombinant plasmids, which is discussed later in this technique primer.


Access options

Get full journal access for 1 year

All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.

Get time limited or full article access on ReadCube.

All prices are NET prices.


Methods requiring DNA recombination in mammalian cells

The classical method to generate E1-substituted adenovirus vectors requires recombination between two DNA molecules, one carrying sequences mapping to the very left end of the adenovirus genome and the gene of interest, and the other carrying sequences that slightly overlap the 3′-most viral sequences on the first molecule and continue to the right end of the adenovirus genome. The latter, which we will refer to as right end sequences for simplicity, can be either a linear DNA purified from virions, or a plasmid. This recombination takes place inside E1-expressing cells such as 293 cells, generating the desired recombinant viral DNA.

Originally, the right end of the adenovirus genome was obtained by digesting viral DNA with a restriction enzyme(s) that cuts in the E1 region (Figure 2a).42 Because of the small difference in size between the undigested and digested viral DNAs, it is difficult to verify the completeness of the digest by agarose gel electrophoresis or to separate the two DNAs for purification. This can be a problem because undigested viral DNA will generate virus more efficiently than the recombination product, leading to the contamination of the recombinant virus preparation with the parental virus. Moreover, the small restriction fragments that correspond to the left end of the genome and are carried over during the transfection can religate inside the cell and generate replication-competent adenovirus.4344 Therefore two or three rounds of purification by plaque assay must be performed, and multiple plaques must be analyzed to identify the recombinant virus. A second disadvantage of this method is that the recombination event, which is required to link the left adenovirus sequences carrying the gene of interest to the right end sequences, is inefficient in mammalian cells. Therefore it usually takes up to 2 weeks for the recombinant virus to appear.

(a) Construction of E1-substituted adenovirus vectors by classical recombination in helper cells. See text for explanations and Figure 1 legend for symbol definitions. The double arrow represents the expression cassette. Hollow and black viruses at the bottom of the panel indicate parental and recombinant viruses, respectively. (b) Construction of E1-substituted adenovirus vectors from infectious circular adenovirus DNAs. Ori, Amp: colE1 origin of replication and ampicillin-resistance gene, respectively. The tables summarize the main characteristics of both techniques and their derivatives. In each table, features that are mentioned more than once are referred to by numbers in parenthesis.

Three methods have been developed to facilitate the detection and purification of the recombinant virus. By using a viral DNA in which the E1 region is replaced by a cassette expressing the HSV thymidine kinase (tk) gene,45 counter-selection against the parental background virus can be performed by adding ganciclovir to the agar overlay. This counter-selection is reliable only on a secondary plaque assay, however, as the primary plaques obtained directly after transfection may originate from cells that have taken up multiple viral genomes and contain both the tk-expressing parental virus and the recombinant virus. Similarly, viral DNA in which the E1 region is replaced by a cassette expressing E. coli β-galactosidase43 or a green fluorescent protein46 can be used as parental DNA. Plaques containing background virus are detected either by staining with X-gal, or by fluorescence microscopy, respectively. These two methods reportedly allow the isolation of the recombinant virus during the first round of purification. It would be more cautious however, and proper virological procedure, to perform a second round of plaque purification, as suggested for the tk-expressing virus.

The adenovirus DNA–terminal protein complex can generate viral plaques with an efficiency two to three orders of magnitude higher than naked DNA prepared with proteinase. For decades, such complexes have been used to construct recombinant adenoviruses.47484950 More recently, a technique has been adapted in which the DNA–terminal protein complex is digested with a restriction enzyme at several sites in the Ad5 genome, better preventing the regeneration of the parental virus.51 The expression cassette is cloned into a cosmid containing a full-length copy of the adenovirus genome. Both the circular cosmid and the digested viral DNA are transfected into helper cells. A few hundred plaques, which arise by recombination of the full-length cosmid-derived genome with the two terminal protein-linked genome ends, are usually obtained using this procedure, among which about 70% are positive for the presence of the expression cassette.

To overcome the inefficiency of homologous recombination in mammalian cells, a technique has been developed that uses the bacteriophage P1 Cre-lox recombination system.52 This system is composed of three elements: (1) a recombinant adenovirus that contains two loxP sites flanking the packaging signal (2) a shuttle vector which contains the left ITR, the packaging signal, the expression cassette and a loxP site (3) a 293-derived cell line which expresses the phage P1 Cre recombinase. To generate E1-substituted adenovirus vectors, the shuttle vector carrying the gene of interest and the viral DNA are co-transfected into the helper cells. First, an intramolecular recombination occurs between the two loxP sites present in the viral DNA. This generates an adenovirus genome which is able to replicate and express viral genes, but is unable to be packaged. A second recombination occurs between the loxP site of this product and that on the shuttle vector, generating the desired recombinant virus. The main disadvantage of this method is the persistence of the parental virus in the preparation, which can represent up to 30% of the viral population after the first passage, and still constitutes 0.2% after three passages. Therefore, this method requires a careful verification of the identity of the recombinant virus.

A very recent improvement of this technique was obtained by using adenovirus DNA isolated from plasmids instead of viral particles.53 A large plasmid was constructed that contains the Ad5 backbone devoid of the left ITR, the packaging signal, and the E1 sequences. This plasmid, linearized at a site flanking the right ITR, is co-transfected into helper cells with a shuttle plasmid that contains the left adenovirus sequences and the expression cassette. Virus is generated upon homologous DNA recombination between both DNA molecules. Unlike the above-described methods, contamination with parental virus is not possible, and therefore a limited number of plaques needs to be analyzed.

Another approach to construct recombinant adenovirus vectors relies on the ability of circular adenovirus DNAs to generate virus upon transfection into 293 cells (Figure 2b).2054 These circular viral DNAs contain an ampicillin resistance gene and a bacterial origin of replication in place of the E1 region, and are therefore able to replicate in E. coli as plasmids. These vectors have been subsequently modified to increase their size to about 40 kb, or have been deleted for their packaging signal. These features render the parental molecules unable to be packaged into virions, and therefore a homogeneous recombinant virus preparation should be obtainable. In this system, the gene of interest is cloned into a small plasmid containing the left end of the adenovirus genome. The resulting plasmid, linearized, is co-transfected into helper cells with the circular adenovirus plasmid. Recombination must occur between both DNAs to create a molecule that can be replicated and packaged to generate the recombinant virus. The same technique can be used to insert expression cassettes in the E3 region, or in both the E1 and E3 regions. However, manipulation of these large plasmids in E. coli is rendered difficult because of the presence of a 200-bp long palindromic sequence that results from the juxtaposition of the left and right adenovirus ITRs in the plasmid construct. Such structures are known to be unstable in E. coli and deleterious for bacterial growth.55 Moreover, the formation of plaques upon transfection of circular adenovirus DNA has been reported to be inefficient,41 probably because imbedded adenovirus ITRs initiate DNA synthesis less efficiently than those at the ends of linear DNA.56 A 293 cell line constitutively expressing the adenovirus preterminal protein and DNA polymerase can be used to overcome this replication problem.41

As for the techniques using linear viral DNAs, this homologous recombination method was improved using the Cre-lox recombination system. In a first embodiment, loxP sites were inserted in both the large circular adenovirus DNA and the small shuttle plasmid.57 Recombinant adenovirus is obtained as a result of Cre-mediated recombination between both plasmids after their cotransfection into a 293 cell line expressing Cre recombinase. The frequency of virus rescue using this specific recombination system is significantly higher (approximately 30-fold) than by in vivo homologous recombination. In a second embodiment, the E1 region of the circular form of adenovirus DNA is replaced with a cosmid vector flanked by loxP sites.58 As above, this replacement produces a DNA molecule too large to be packaged into virions. Expression cassettes are inserted between the loxP-flanked cosmid backbone and the adenovirus genome via packaging into phage λ. The resulting cosmid is transfected into helper cells that express the Cre recombinase. An intramolecular Cre-lox mediated recombination excises the cosmid vector backbone and produces a molecule that can be packaged into virions. Using this method, viral plaques are observed between 8 and 10 days after transfection.


Results

Efficient inactivation of pigment gene olvA in A. niger by CRISPR/Cas9 system embodied HH-sgRNA-HDV element

To test the feasibility and efficiency of the CRISPR/Cas9 system in A. niger, we choose to target the olvA gene which can be easily screened phenotypically after inactivation. olvA gene encodes the YWA1 hydrolase homolog and its deletion can form olive conidia on regeneration medium while wild strain shows black conidia. A 20 bp of the protospacer sequence with the 3′-PAM AGG for olvA was chosen at the start of the ORF (nucleotide − 18 to − 37). The resulting plasmids pAN1 was transformed into A. niger through protoplast transformation. Most transformants (24/26) grown on hygromycin plates (Fig. 4a) showed the desired phenotype of olive colonies, with the mutational rate of over 90%. Because conidia of A. niger contain two or more nuclei, the isolation of single mutant colony from the original transformants is necessary. We diluted the conidia of primary transformants to 10 4 –10 5 times and streak the conidia diluent on medium with hygromycin for growth of 3 days to isolate the homozygotes (Fig. 4b). Afterwards, the genome DNA extracted from the single positive colony was used as the template for PCR and DNA identification. The sequencing results reflected either single nucleotide deletion or mutation in upstream of the PAM motif in olvA gene in the two transformants (Table 2).

Primary and the second generation of transformants in which olvA was inactivated by CRISPR/Cas9 system. a Growth of wild-type strain (WT) and ΔolvA mutants. b Growth of single colony propagated from the primary transformants

Over-expression of glaA and down-regulation of agdF in one step by CRISPR

After transformed by plasmid pAN-agdF and repair fragment containing glaA expression cassette and 1 kb arms homologous to agdF, A. niger colonies with hygromycin resistance were obtained. PCR verification of the up and down parts of the insertion (Fig. 3b) in the genome of positive transformants was performed. The result shows (Fig. 3c) three out of five candidates possessing the insertion fragment. One of the three was selected as the agdF::glaA mutant for further enzyme activity assay. As it shows in Fig. 5, the activity of glucoamylase of agdF::glaA mutant was 25.9% higher than wild strain, while the activity of α-glucosidase of agdF::glaA mutant was 61.4% lower than wild strain.

The comparison of enzyme activity between wild strain and agdF::glaA mutant. a Comparison of the activity of glucoamylase. b Comparison of the activity of α-glucosidase (n = 5, *p < 0.05)

Efficiency of gene replacement with different length of the homologous arm

To test the relation between the efficiency of gene replacement and the length of the homologous arms, we further designed the repair fragments with 2 sets of different arm lengths (500 and 100 bp) based on the above-mentioned experiment. We co-transformed the plasmid pAN-agdF and the repair fragments with different length of homologous arms. After co-transformation, we selected 3 or 5 transformants from each transformation and extracted their genome DNA for PCR identification. And we found all tested transformants (3/3 and 5/5, respectively) were positive. The results of PCR (Fig. 6) demonstrated that a pair of 100-bp homology arms is sufficient to obtain homologous integration stimulated by the CRISPR/Cas9 system in A. niger with high efficiency.

Verification of genotype by PCR in the selected transformants after co-transformation of repair fragments with different lengths of homologous arms. The expected length for amplified product from agdF::glaA mutants was about 820 bp for both up- and down-stream PCR, while no band will be produced from the wide type strain CBS513.88. a PCR results using homologous arm of 500 bp. Results showed that all 3 tested candidates had the expected genotype. b PCR results using homologous arm of 100 bp. Results showed that all 5 tested candidates had the expected genotype


MATERIALS AND METHODS

Cell culture

The human embryonic kidney (HEK293) cells (Invitrogen) were cultured in DMEM supplemented with 10% fetal calf serum (FCS). The HEK293 cells constitutively expressed Adeasy deleted E1 gene in-trans. Infective virus particles were produced after these cells were transfected with E1-deleted adenovirus vectors. After infection with the recombinant adenoviruses, the cells were maintained in DMEM supplemented with 5% FCS.

Bacteria, plasmids and viral DNA

E.coli strains DH10β and BUN21 and the plasmid pML300 were kindly donated by Prof. Stephen J Elledge from the Harvard Medical School in Boston, (MA, USA) ( 18 ). DH10β was used for generating the recombinant donor plasmid and BUN21 was used for generating and propagating the recombinant adenoviral plasmids. The plasmid pML300 contained in BUN21 carries the red and gam recombinase gene induced by rhamnose, and is unable to replicate when the bacteria are grown at 42°C ( 18 ). The plasmid MAGIC1 and the plasmid 1202 were also provided by Prof. Stephen J Elledge ( 18 ). The adenovirus bone vector (pAdeasy) and pShuttle plasmid were obtained from Stratagene ( 15 ). We constructed the novel donor plasmid pRTRA ( Figure 1 ), the recipient plasmid pAd-pheS (full-length adenoviral genome), the plasmid pPic-man and the plasmid pShuttle-cmv-red-sv40polA.

Cloning the foreign genes gfp and man into the donor plasmid using restriction enzyme Bsu36I and T4 DNA polymerase

The gfp gene was amplified from pEGFP-1 (Clontech) by PCR. The forward primer was 5′- TTAC GATGGTGAGCAAGGGCGAGGA-3′, and the reverse primer was 5′- TGAC TTACTTGTACAGCTCGTCCATGCC-3′. The man gene was amplified from pPic-man using the primers: ZL15F, 5′- TTACT GAAGCGCATACTGTGTCGCC-3′ and ZL16R, 5′- TGAC CGGATTCACTCAACGATTGG-3′. The amplified fragments were incubated with 0.5 U of T4 DNA polymerase and 4 mM dGTP (TaKaRa) at 12°C for 45 min, as described previously ( 19 , 20 ). For each gene, a total of 30 ng of treated fragments and 1 ng of the Bsu36I-digested ( CCTTAGG and CCTGAGG ) pRTRA were ligated with 5 U of T4 DNA ligase™ (TaKaRa) at 16°C for 12 h in 1 μl buffer. The gfp gene and man gene were inserted into pRTRA to form the plasmids pRTGA and pRTMA separately ( Figure 2 ).

Modification of the donor plasmid

The fragment of chloramphenicol resistent gene was amplified from pBT (Strategene) by PCR using the forward primer [5′-TTT GTCGACATAACTTCGTATAATGTATGCTATACGAAGTTAT ACGGGGAGAGCCTGAGCAAAC (SalI and 34 bp loxP sites underlined)] and the reverse primer [5′-TTT GTGCACATAACTTCGTATAGCATACATTATACGAAGTTAT CAGCATCACCCGACGCACTTT-3′ (ApaLI, 34 bp loxP sites underlined)]. The PCR was performed at 95°C for 5 min, followed by 25 cycles at 95°C for 45 s, 56°C for 45 s and 72°C for 1 min by using Thermo Hybrid PX2 (Thermo) and TaKaRa Extaq™ polymerase (TaKaRa). The fragment cut by ApaLI and SalI was ligated into the pRTRA vector (also cut by the same enzymes), resulting in the recombinant plasmid pRTRC.

Generating the recombinant adenoviral vector by MAGIC

The overall strategy developed is shown in Figure 3 . E.coli DH10β containing the donor vector pRTRA was grown in Luria–Bertani (LB) broth containing ampicillin (100 μg/ml).The recipient strain BUN21 containing the plasmid pML300 and the recipient plasmid pAd-pheS was grown in LB broth containing spectinomycin (50 μg/ml), kanamycin (50 μg/ml) and glucose (0.2% w/v) overnight. The recipient strain was washed twice with 2 volume of LB broth the next day. The donor and recipient strains were separately diluted to 1:25, 1:50, 1:100 or 1:200 with LB broth containing 0.2% w/v rhamnose and grown at 30°C for 2 h to an A 600 of 0.15∼0.25, and the donor and recipient strains were mixed to a ratio of 1:1 based on their A 600 in the presence of 0.2% w/v l -arabinose. The mixture was incubated at 37°C for 2 h without shaking, and then for a further 2 h with shaking. The recombinant culture was diluted at the ratio of 1:100, plated on the selective plates containing kanamycin (50 μg/ml), ampicillin (100 μg/ml), 10 mM Cl-Phe and 0.2% w/v l -arabinose, and finally incubated at 42°C overnight ( 18 ).

Production of recombinant adenoviruses and proliferation

The recombinant adenoviral plasmids were amplified by incubating the colony identified in 50 ml LB broth containing kanamycin (50 μg/ml), ampicillin (100 μg/ml) and 0.2% w/v arabinose. The culture was grown overnight at 37°C. The maxiprep DNA was then prepared from the liquid culture and digested in a sufficient amount (5 μg of DNA) with PacI. Subsequently, the buffer and the excess enzyme from restriction reactions were removed by phenol extraction/ethanol precipitation. The DNA was re-suspended in 50 μl of sterile 0.1× TE buffer or dH 2 O, and then added to HEK293 cells in the presence of Lipofectamine 2000 (Invitrogen). At 4 h post-transfection, the cells were incubated in 5 ml DMEM containing 5% FCS. About 8 days after transfection, the cells were harvested and pelleted by low-speed centrifugation, and the viruses were liberated by three freeze/thaw cycles. The cell lysate [1 ml in 1× phosphate-buffered saline (PBS) (pH 7.4)] containing the recombinant adenoviruses was then amplified and purified. To generate higher titer viral stocks, the HEK 293 cells were infected by the cell lysate and the harvest process was repeated.

Construction of a model adenoviral cDNA expression library

The donor strain containing pRTRA and the donor strain containing pRTGA were mixed at ratios of 1:30, 1:300, 1:3000 and 1:30 000 on the basis of their A 600 . The mixed donor stains at different ratios were mixed with the recipient strain at a 1:1 ratio on the basis of their A 600 ( 3 , 18 ). The subsequent experiments were based on the complete MAGIC procedure as reported previously ( 18 ). LB broth was used to wash all the colonies on the selective plates which were then incubated overnight in 100 ml LB broth containing 50 μg/ml kanamycin, 100 μg/ml ampicillin, 0.2% w/v l -arabinose and 10 mM Cl-Phe. The DNA was extracted for greater yields, treated with PacI, and the excess buffer and enzyme were removed by phenol extraction/ethanol precipitation. The DNA (5 μg) was re-suspended in 100 μl of sterile 0.1× TE buffer or dH 2 O, and then added to a 60 mm plate containing 1 × 10 6 HEK293 cells that had been incubated in DMEM with 5% FCS for 2 h. At 4 h post-transfection, the cells were incubated for 8 days in 5 ml DMEM containing 5% FCS. After three freeze-thaw cycles, the resultant lysate (1 ml) was used to infect HEK 293 cells in a 10 cm plate (70% confluent) for 1 h before being incubated with 5 ml DMEM supplemented with 5% FCS.


The logic of selfish genetic elements

Though selfish genetic elements show a remarkable diversity in the way they promote their own transmission, some generalizations about their biology can be made. In a classic 2001 review, Gregory D.D. Hurst and John H. Werren proposed two ‘rules’ of selfish genetic elements.[4]

Rule 1: The spread of selfish genetic elements requires sex and outbreeding

Sexual reproduction involves the mixing of genes from two individuals. According to Mendel’s Law of Segregation, alleles in a sexually reproducing organism have a 50% chance of being passed from parent to offspring. Meiosis is therefore sometimes referred to as �ir”.[30]

Highly self-fertilizing or asexual genomes are expected to experience less conflict between selfish genetic elements and the rest of the host genome than outcrossing sexual genomes.[31�] There are several reasons for this. First, sex and outcrossing put selfish genetic elements into new genetic lineages. In contrast, in a highly selfing or asexual lineage, any selfish genetic element is essentially stuck in that lineage, which should increase variation in fitness among individuals. The increased variation should result in stronger purifying selection in selfers/asexuals, as a lineage without the selfish genetic elements should out-compete a lineage with the selfish genetic element. Second, the increased homozygosity in selfers removes the opportunity for competition among homologous alleles. Third, theoretical work has shown that the greater linkage disequilibrium in selfing compared to outcrossing genomes may in some, albeit rather limited, cases cause selection for reduced transposition rates.[34] Overall, this reasoning leads to the prediction that asexuals/selfers should experience a lower load of selfish genetic elements. One caveat to this is that the evolution of selfing is associated with a reduction in the effective population size.[35] A reduction in the effective population size should reduce the efficacy of selection and therefore leads to the opposite prediction: higher accumulation of selfish genetic elements in selfers relative to outcrossers. Empirical evidence for the importance of sex and outcrossing comes from a variety of selfish genetic elements, including transposable elements,[36,37] self-promoting plasmids,[38] and B chromosomes.[39]

Rule 2: The presence of selfish genetic elements is often revealed in hybrids

The presence of selfish genetic elements can be difficult to detect in natural populations. Instead, their phenotypic consequences often become apparent in hybrids. The first reasons for this is that some selfish genetic elements rapidly sweep to fixation, and the phenotypic effects will therefore not be segregating the in the population. Hybridization events, however, will produce offspring with and without the selfish genetic elements and so reveal their presence. The second reason is that host genomes have evolved mechanisms to suppress the activity of the selfish genetic elements, for example the small RNA administered silencing of transposable elements.[40] The co-evolution between selfish genetic elements and their suppressors can be rapid, and follow a Red Queen dynamics, which may mask the presence of selfish genetic elements in a population. Hybrid offspring, on the other hand, may inherit a given selfish genetic element, but not the corresponding suppressor and so reveal the phenotypic effect of the selfish genetic element.[41,42]


Restriction map & virtual digest

DNA maps are simplified representations of nucleotide sequences, showing features of particular interest such as genes. Restriction maps are DNA maps that show restriction sites, the locations where specific restriction enzymes will cut the DNA. Here is a map of pGLO:

I made this map with Benchling, the online software package that I usually use for analyzing nucleotide sequences. Some of the features on this pGLO map should look familiar, such as the ampicillin resistance gene (AmpR) and the GFP gene. Also included are restriction sites for three enzymes that we have in lab: HindIII, PvuI, and EcoRI. This map will give you a general idea of what size DNA fragments to expect if you cut pGLO with each of these enzymes:

  • PvuI: Cuts once, so you get one linear piece, the size of the entire plasmid (5371 bp).
  • EcoRI: Cuts once, so you get one linear piece, the size of the entire plasmid (5371 bp). Although EcoRI cuts in a different place than PvuI, the resulting fragment will be the same size.
  • HindIII: Cuts twice, so you will get a small piece and a large piece.

Now suppose you want to cut the plasmid with more than one enzyme at a time. Each enzyme will act on its own restriction sites, so the different enzymes won't affect each other. From the map above, you can see that if you cut with both EcoRI and HindIII, you will cut out a tiny piece between the EcoRI and HindIII sites you probably wouldn't be able to see this small fragment on a gel, so it's not worthwhile to perform this digest. Suppose you cut with PvuI and EcoRI together. Using this slightly more detailed map (courtesy of Snapgene), you should be able to predict the exact sizes of the DNA fragments you will get:

This map shows the exact locations of the restriction sites. Each nucleotide is assigned a number, based on an arbitrarily chosen starting point. If PvuI cuts at nucleotide 3054 and EcoRI cuts at nucleotide 2063, and the entire circular plasmid is 5371 bp long, you can probably figure out the sizes of the fragments you will get.

Unfortunately, the map above leaves out the HindIII sites (I don't know why they weren't included).

Perform some virtual digests of pGLO

Using the restriction enzymes we have in lab, the best combination for getting nicely spaced fragments of pGLO on your gel would be HindIII and PvuI. To figure out the expected results of the HindIII + PvuI digest, you can use restriction mapping software such as this one:

For a precise electronic restriction digest, you need to enter the nucleotide sequence of the DNA you're going to cut. Find the pGLO sequence (the original sequence from Bio-Rad) on the Sequence Data page (keep that page open in another tab you'll use it again).

Copy the whole sequence, including the numbers, and paste it into the "sequence info" box in the Restriction Analyzer (the software will ignore the numbers and spaces, and only look at the nucleotide sequence). Note that the sequence is shown for only one strand of DNA, but the software is smart enough to recognize that the actual DNA is double-stranded. Check "Circular" under Conformation, and then go to the Include box and select PvuI. Click on Virtual Digest.

Virtual digest will take you to a page showing the complete nucleotide sequences of the DNA fragments produced in the digest (which you probably don't care about) and also the length of each fragment (which you will be able to verify on your gel. Those fragment lengths are your expected results for this experiment.

Repeat the virtual digest procedure for the enzyme SspI.

Alternative approach: You could also use Benchling instead of RestrictionMapper. Benchling is more modern and more powerful, but it takes a little while to learn and you will have to sign up for a (free) account. Personally, I use Benchling for most of my sequence work, and I recommend it.

Make your own map of pGLO

Draw your own simplified map of pGLO. Show the exact cut sites for PvuI and SspI. Also include the approximate locations of the GFP, AmpR (ampicillin resistance, also called β-lactamase or bla), and AraC genes.

Perform virtual digests of pGLO non-fluorescent mutant

The Bio 6B Special Projects students accidentally created a mutant version of pGLO that doesn't have a functional GFP gene. Find the sequence for the pGLO Non-Fluorescent Mutant on the sequence page.

Perform virtual digests of this sequence the same way you did with the original pGLO sequence. You should get a slightly different result for one of the enzymes.

On the pGLO map, show where the mutant sequence differs from the original.

Perform a virtual digest of lambda

Lambda bacteriophage is the DNA you'll use as a molecular weight marker. This DNA is widely available gets cut into a range of predictably sized fragments, making it one of the most common molecular weight markers.

Find the sequence of lambda on the Sequence Data page. The lambda genome is linear DNA, unlike the circular plasmids.

Perform a virtual digest of lambda with the enzyme Hind III and write down the fragment sizes. Unlike the plasmids, lambda DNA is linear, so check the appropriate box before doing the digest.

Diagram the gel results

Draw a labeled picture of the expected gel results for the following restriction digests:

  • pGLO Green cut with PvuI
  • pGLO Green cut with SspI
  • pGLO Non-Fluorescent cut with PvuI
  • pGLO Non-Fluorescent cut with SspI
  • Lambda DNA cut with Hind III

Draw a diagram of the gel showing the relative positions of the bands, and label each band with its size in base pairs. This will represent your expected results for the actual digest that you perform.

How much DNA per band?

If you perform a restriction digest and run it on a gel, you'd like to be able to see all the bands. Your ability to see a band on the gel depends on how many nanograms of DNA are in that band, as well as how you stain the DNA. We use ethidium bromide in our gels, which allows us to visualize individual bands containing as little as 50 ng. However, the different bands from a digest won't have the same mass of DNA. If you cut some DNA into two fragments and one fragment is twice as large as the other, the larger fragment will have twice the mass of the smaller fragment.

Suppose you are doing a restriction digest of lambda using the enzyme Hind III. If you start with 500 ng of uncut lambda DNA, how many nanograms of DNA will be in each of the bands on your gel? Label the bands in the Lambda/Hind III lane of your gel diagram with the amount in nanograms. Do you think you'll be able to see the smallest band?

What to turn in

Working with your lab partners, make one pGLO map (showing cut sites for both the unmutated and mutated versions) and one gel picture for your lab group. Label the gel diagram in terms of base pairs and (for Lambda/Hind III only) nanograms. Turn the diagrams in today in lab, with the names of everyone who worked on it. This counts as a quiz.


Gene (genome) editing to produce genetically modified animals with gene knock-out or knock-in

The initial methods used to generate transgenic livestock resulted in random transgene insertion [10] therefore, new technologies are needed to enable better gene targeting with a higher efficiency in livestock. Although it is conceptually simple to deliver DNA into a fertilized egg via pronuclei injection, this method is technically challenging and the injected DNA construct is integrated randomly into the genome, resulting in unpredictable transgene expression profiles. In addition, microinjection can damage the zygote and requires expensive equipment. These short-comings can be partly overcome by the development of gene (genome) editing approach, which uses a designer nuclease (as a pair of molecular scissors) to generate a double strand break (DSB) in DNA at a desired genomic locus (Fig. 5). Thereafter, one of two endogenous repair mechanisms may repair the DSB DNA: non-homologous end joining (NHEJ) and the homology-directed repair (HDR) [38]. In the error-prone NHEJ pathway, the two ends of the DSB DNA are brought together and ligated without a homologous template for repair, which often inserts or deletes nucleotides (indels). If an indel results in a frame shift mutation, the target gene may lose function (knockout). The HDR pathway requires the provision of an exogenous DNA template along with a site-specific genome editing nuclease to repair the DSB DNA, thereby causing the knock-in of a desired sequence of DNA into the genome of an embryo or animal cells [67]. In practice, modification of a targeted gene is commonly achieved by microinjecting, into an embryo obtained by in vitro fertilization or intracytoplasmic transfer, a gene editing system that consists of a guide RNA and the Cas9 endonuclease, and, if necessary, a repair DNA template [68]. The guide RNA provides sequence specificity to target the Cas9 endonuclease to a complementary site in the genome for creating a DSB.

Gene (genome) editing of animals using the ZFN, TALEN or CRISPR/Cas9 technique. a designer nuclease (ZFN, TALEN or CRISPR/Cas9) cleaves a DNA molecule to generate a double strand break (DSB) at a desired genomic locus. Thereafter, one of two endogenous repair mechanisms may repair the DSB DNA: non-homologous end joining (NHEJ) and the homology-directed repair (HDR). In the NHEJ pathway, the two ends of the DSB DNA are brought together and ligated without a homologous template for repair, which often inserts or deletes nucleotides (indels) to cause gene disruption (knockout). The HDR pathway requires the provision of an exogenous DNA template along with a site-specific genome editing nuclease to repair the DSB DNA, thereby causing the knock-in of a desired sequence of DNA into the genome of an embryo or animal cells. Because of its more precise targeting of genes, CRISPR/Cas9 is gaining momentum in life sciences as the preferred editor of gene editing of livestock species

An earlier designer nuclease was zinc finger nuclease (ZFN the first gene editing tool), and a subsequently discovered designer nuclease is transcription activator-like effector nuclease (TALEN, the second gene editing tool), both of which are modular proteins containing an adaptable DNA-binding domain. The ZFN method involves engineering a protein that contains both a zinc finger DNA-binding domain and a restriction endonuclease domain. The TALEN approach utilizes engineered enzymes containing a DNA-binding domain and a separate DNA-cleaving domain. In recent years, CRISPR (clustered regularly interspaced short palindromic repeats)-associated nuclease-9 (CRISPR/Cas9) has been used as a designer nuclease to provide a more efficient, more accurate, more versatile, more robust, and simpler tool in genomic engineering [19, 62]. ZFN, TALEN, or CRISPR/Cas9 components are delivered into target cells through transfection (lipid-based agents, electroporation, nucleofection, or microinjection) or bacteriophages, depending on cell type and plasmid [69,70,71].

TALENs and CRISPR/Cas9 were first successfully used in pigs in 2013 [72] and 2014 [59, 73], respectively. Over the past 5 years, CRISPR/Cas9 has rapidly gained momentum as the favored gene editor for livestock species. The CRISPR-Cas9 system was discovered in 2007 in bacteria (e.g., a genus of gram-positive cocci or spherical bacteria) and archaea, and is used naturally to defend against invading viruses (bacteriophages). In response to a viral infection, the bacterial CRISPR/Cas9 is guided by a short RNA fragment known as a guide RNA to snip off a piece of viral DNA, creating a DSB in its target loci [68]. The guide RNA is complementary to a segment of the genome of the targeted organism, so that the Cas9 nuclease will cleave DNA with a high degree of specificity. Of note, recognition of the target DNA by Cas9 is dependent on the presence of a short protospacer adjacent motif (PAM) sequence located directly downstream on the untargeted DNA strand [38]. Thus, the CRISPR system consists of two components (a Cas9 endonuclease and a guide RNA) as a ribonucleoprotein. Experimentally, the guide RNA can be designed using molecular biology tools in the laboratory to direct Cas9 to a specific DNA sequence for cleavage at virtually any genomic locus. The milestones for the use of CRISPR/Cas9 in producing gene-edited swine are shown in Table 2.

Advantages and applications

Traditional livestock breeding is beset with such problems as long breeding cycles and limitations of genetic resources. In contrast, genome editing tools can provide more precise, more specific, more predictable and more rapid solutions to solving these problems at relatively affordable costs [38]. Thus, besides knocking out gene function, CRISPR can be employed to delete large DNA fragments from the genome of an animal. Furthermore, a gene editing technique requires fewer steps and has a higher efficiency than the previous methods of animal transgenesis. For example, studies with livestock zygotes have shown a 30% editing frequency with ZFN, TALEN and CRISPR/ Cas9 techniques [19, 72,73,74,75,76,77]. Compared to other gene silencing techniques such as RNAi and antisense RNA, CRISPR/Cas9 offers a higher efficiency, an ability to cleave methylated loci, greater ease of design, and greater flexibility [68]. It should be borne in mind that knockout of a gene provides a cleaner phenotype than its knockdown and that production of knockout pigs does not necessarily require application of a genome editing system. Several laboratories have reported success with producing gene-edited pigs, which can potentially serve as organ donors, disease models, bioreactors, inactivation of porcine endogenous retrovirus in pigs, or founder animals of genetic lines with enhanced productivity (e.g., muscle growth) or disease resistance traits [59,60,61,62,63,64]. Thus, gene editing increases successes in single-gene and multi-allelic modifications of the livestock genome, as well as in site-specific introductions of foreign genes during embryogenesis.

There are many examples for the genome editing-based production of transgenic pigs with important production and disease-resistance traits, including nutrient utilization and meat production as well as resistance to viral infections and metabolic disorders (Table 3). First, disruption of the myostatin (a negative regulator of myogenesis) gene using TALEN as an editor successfully created myostatin-knockout pigs, which exhibited a double-muscled phenotype, greater body weight, greater longissimus muscle mass, and a 100% increase in the number of muscle fibers than wild-type pigs [78]. Second, utilizing the CRISPER/Cas9 technology, Zheng et al. [79] produced pigs with a functional uncoupling protein 1 (UCP1). UCP1 is expressed in the brown adipose tissue of many animal species and is responsible for nonshivering thermogenesis, thereby playing a crucial role in protecting against cold and regulating energy homeostasis. However, modern pigs lack functional UCP1 genes and are therefore susceptible to cold stress, resulting in a high rate of neonatal mortality, and also spontaneous accumulation of a large amount of white adipose tissue in the body, leading to reduced production performance [3]. Of note, insertion of the mouse adiponectin-UCP1 gene into the porcine endogenous UCP1 locus via the CRISPR/Cas9 as an editor can generate UCP1-knockin pigs that exhibit an improved ability to maintain body temperature, a decreased white fat mass, and an increased lean carcass yield [79]. Third, CRISPR/Cas9 gene targeting and SCNT technologies have been used to create pigs without the CD163 gene that encodes a cellular receptor for the porcine reproductive and respiratory syndrome virus-1 (PRRSV-1, also referred to as “blue ear disease” virus) [59]. Whitworth et al. [60] reported that pigs with the CD163 knock-out were fully resistant to the PRRSV-1 (European strain) and PRRSV-2 (North American strain). Similar results were observed by Burkard et al. [63] against both PRRSV-1 and PRRSV-2, and by Yang et al. [64] against a highly pathogenic PRRSV strain (belonging to the North American strain) isolated in South China. Interestingly, Wells et al. [61] found that genetically modified pigs, which were produced through the replacement of porcine CD163 scavenger receptor cysteine-rich domain 5 with a CD163-like homolog, were resistant to PRRSV-1 but not to PRRSV-2. Males and females can be used as breeding stocks to produce generations of PRRSV-resistant offspring.

Disadvantages

Although the ZFN method provided the first breakthrough in site-specific gene editing, it has some limitations, such as off-target cutting of the DNA, cytotoxicity, expensive, time-consuming, low efficiency (thus only one genomic edit at a time), and technical challenges to prepare effective ZFN tools [38]. Compared with the ZFN editor, the TALEN technique is more flexible in genetic engineering because its DNA-binding domain can target a wider range of DNA sequences. Although the TALEN editor is easier to design than the ZFN, the TALEN method is expensive and technically difficult when the goal is to simultaneously make multiple edits to the genome [68]. In addition, delivering the gene-editing Cas9 directly to embryos by microinjection remains a challenging process, and microinjection itself may damage the embryos. Compared to the ZFN and TALEN, CRISPR/Cas9 is known to have a higher frequency of off-target effects [80]. Another major obstacle to the use of the CRISPR/Cas9 technology for generating gene-edited animals is the problem of mosaicism (the presence of more than one genotype in one individual) that is common in founder animals [68]. Furthermore, for all currently available gene-editing methods, the rates of prenatal mortality in gene-edited fetuses are much greater than those for control fetuses. To date, the efficiency of gene editing in livestock, including swine, remains suboptimal. The procedures for gene editing should be easier and cheaper, so that more producers can utilize this innovative technique on their own farms for improving animal breeding.


ACKNOWLEDGMENTS

This work was supported by the Slovenian Research Agency under grant agreement no.(Z2-7257) and was in part carried out at the Faculty of Health Sciences, University of Primorska, Izola, Slovenia. I kindly thank Maria Pilar Garcillán-Barcia, Fernando de la Cruz (UNICAN, Spain), and Joshua Ramsey (Curtin Univ., Australia) for sharing and discussing data, Filip Buric (Chalmers Univ. of Tech., Sweden) for commenting on the manuscript, and Tomaz Pisanski (UP-FAMNIT, Slovenia) and Ales Lapanje (IJS, Slovenia) for helpful discussions in their respective fields of research.