LKB1 preserves genome integrity by stimulating BRCA1 expression.

Serine/threonine kinase 11 (STK11, also known as LKB1) functions as a tumor suppressor in many human cancers. However, paradoxically loss of LKB1 in mouse embryonic fibroblast results in resistance to oncogene-induced transformation. Therefore, it is unclear why loss of LKB1 leads to increased predisposition to develop a wide variety of cancers. Here, we show that LKB1 protects cells from genotoxic stress. Cells lacking LKB1 display increased sensitivity to irradiation, accumulates more DNA double-strand breaks, display defective homology-directed DNA repair (HDR) and exhibit increased mutation rate, compared with that of LKB1-expressing cells. Conversely, the ectopic expression of LKB1 in cells lacking LKB1 protects them against genotoxic stress-induced DNA damage and prevents the accumulation of mutations. We find that LKB1 post-transcriptionally stimulates HDR gene BRCA1 expression by inhibiting the cytoplasmic localization of the RNA-binding protein, HU antigen R, in an AMP kinase-dependent manner and stabilizes BRCA1 mRNA. Cells lacking BRCA1 similar to the cell lacking LKB1 display increased genomic instability and ectopic expression of BRCA1 rescues LKB1 loss-induced sensitivity to genotoxic stress. Collectively, our results demonstrate that LKB1 is a crucial regulator of genome integrity and reveal a novel mechanism for LKB1-mediated tumor suppression with direct therapeutic implications for cancer prevention.

Direct Measurement of B Lymphocyte Gene Expression Biomarkers in Peripheral Blood Transcriptomics Enables Early Prediction of Vaccine Seroconversion.

Peripheral blood transcriptome is a highly promising area for biomarker development. However, transcript abundances (TA) in these cell mixture samples are confounded by proportions of the component leukocyte subpopulations. This poses a challenge to clinical applications, as the cell of origin of any change in TA is not known without prior cell separation procedure. We developed a framework to develop a cell-type informative TA biomarkers which enable determination of TA of a single cell-type (B lymphocytes) directly in cell mixture samples of peripheral blood (e.g., peripheral blood mononuclear cells, PBMC) without the need for subpopulation separation. It is applicable to a panel of genes called B cell informative genes. Then a ratio of two B cell informative genes (a target gene and a stably expressed reference gene) obtained in PBMC was used as a new biomarker to represent the target gene expression in purified B lymphocytes. This approach, which eliminates the tedious procedure of cell separation and directly determines TA of a leukocyte subpopulation in peripheral blood samples, is called the Direct LS-TA method. This method is applied to gene expression datasets collected in influenza vaccination trials as early predictive biomarkers of seroconversion. By using TNFRSF17 or TXNDC5 as the target genes and TNFRSF13C or FCRLA as the reference genes, the Direct LS-TA B cell biomarkers were determined directly in the PBMC transcriptome data and were highly correlated with TA of the corresponding target genes in purified B lymphocytes. Vaccination responders had almost a 2-fold higher Direct LS-TA biomarker level of TNFRSF17 (log 2 SMD = 0.84, 95% CI = 0.47-1.21) on day 7 after vaccination. The sensitivity of these Direct LS-TA biomarkers in the prediction of seroconversion was greater than 0.7 and area-under curves (AUC) were over 0.8 in many datasets. In this paper, we report a straightforward approach to directly estimate B lymphocyte gene expression in PBMC, which could be used in a routine clinical setting. Moreover, the method enables the practice of precision medicine in the prediction of vaccination response. More importantly, seroconversion could now be predicted as early as day 7. As the acquired immunology pathway is common to vaccination against influenza and COVID-19, these biomarkers could also be useful to predict seroconversion for the new COVID-19 vaccines.

Oligonucleotide-Directed Mutagenesis by Elimination of a Unique Restriction Site (USE Mutagenesis).

In this method, two oligonucleotide primers are hybridized to the same strand of a denatured double-stranded recombinant plasmid. One primer (the mutagenic primer) introduces the desired mutation into the target sequences, whereas the second primer carries a mutation that destroys a unique restriction site (called "unique site elimination," or USE) in the plasmid. The product of the first part of the method is a heteroduplex plasmid consisting of a wild-type parental strand and a new full-length strand that carries the desired mutation but no longer contains the unique restriction site. In the second phase of the method, the mixed population is incubated with the restriction enzyme that cleaves the unique site. The wild-type molecules are linearized, and the mutated plasmids are resistant to digestion. The mixture of circular heteroduplex DNA and linear wild-type DNA is then used to transform a strain of Escherichia coli that is deficient in repair of mismatched bases. The circular heteroduplex molecules begin to replicate while many of the linear wild-type molecules are unable to reestablish themselves in E. coli cells. Because the mismatched bases are not repaired, the first round of replication generates a wild-type plasmid that carries the original restriction site and a mutated plasmid that does not. DNA from the first set of transformants is recovered, digested once more with the same restriction enzyme to linearize the wild-type molecules, and then used to transform a standard laboratory strain of E. coli.

Saturation Mutagenesis by Codon Cassette Insertion.

Saturation mutagenesis by cassette insertion introduces a library of site-specific changes into a specific DNA sequence within a target gene and is especially useful for analyzing the effect of specific residues on the structure and function of a protein. In this protocol, a set of 11 universal oligodeoxyribonucleotide cassettes is used to generate mutations. The major advantage of this method is that a single set of mutagenic codon cassettes can be used to insert codons encoding all possible amino acids at any predetermined site within a gene. Each of the 11 cassettes contains two recognition sequences for SapI, a restriction enzyme that cleaves outside of its recognition sequence. The recognition sequences for SapI are arranged in opposite orientations and are separated by a central spacer. At the end of each cassette is a 3-bp direct repeat, positioned such that the sites of SapI cleavage bracket each repeat. Cleavage by SapI will result in the generation of three base-cohesive single-stranded ends on the end of the cassette. These three base-cohesive single-stranded ends can then be ligated together to regenerate the original 3-bp direct repeat, while excising the central spacer. It is this 3-bp repeat sequence that is ultimately incorporated into the template. By substituting the 3-bp direct repeats in the universal cassette with other sequences, one can essentially generate all possible amino acid substitutions.

Random Scanning Mutagenesis.

In vitro oligonucleotide and polymerase chain reaction (PCR)-based mutagenesis is generally used for altering the nucleotide sequence of genes to study their functional importance and the products they encode. A thorough approach to this problem is to systematically change each successive amino acid residue in the protein to alanine (i.e., alanine-scanning mutagenesis) or to a limited number of alternative amino acids. Although these strategies can provide useful information, it is sometimes desirable to test a broader spectrum of amino acid changes at the targeted positions. "Random scanning mutagenesis" was developed to examine the functional importance of individual amino acid residues in the conserved structural motif of human immunodeficiency virus (HIV) reverse transcriptase, and this protocol is adapted from that method. This strategy is an oligonucleotide-based method for generating all 19 possible replacements at individual amino acid sites within a protein.

Megaprimer Polymerase Chain Reaction (PCR)-Based Mutagenesis.

The megaprimer method is a simple and versatile approach that can be adopted to create a single mutation in a specific target region as well as to create site-specific insertions, deletions, and gene fusions. This method uses three oligonucleotide primers, two rounds of polymerase chain reaction (PCR), and a DNA template containing the gene to be mutated. The first round of PCR generates a fragment with the desired mutation introduced by using one of the flanking primers and the mutant primer. This amplified fragment-the megaprimer-is used in the second PCR along with the remaining external primer to amplify a longer region of the template DNA. The final product is purified and can be cloned into an appropriate vector. By designing flanking primers with universal restriction site sequences, compatible with the vector of choice, it is possible to create different mutant clones by changing only the mutant primer. Recently, this approach has been improved by the use of forward and reverse flanking primers with significantly different melting temperatures. This allows researchers to perform both PCR steps in a single tube. This protocol has been successfully applied on templates with either low or high G + C content to amplify megaprimers 71-800 bp in length and final products ranging from 400 to 2500 bp.

Multisite-Directed Mutagenesis.

This protocol, suitable for both general and close-proximity mutagenesis, includes a simple and rapid procedure that combines polymerase chain reaction (PCR), DpnI digestion, and overlap extension. The key point of this approach is the use of overlap extension to form a circular DNA plasmid with mutations without the need for phosphorylated primers or ligase reactions. Essentially, during the first round of PCR, the new DNA is synthesized with nicks between the 3' ends of the synthesized DNA and the 5' ends of the first pair of primers. During successive rounds of PCR, a new pair of mutagenic oligonucleotides leads to the synthesis of two DNA segments that anneal together with the overlap sequence inside the two primers. This new mutated molecule also contains nicks but at different positions compared with those formed in the first round of PCR that had been "repaired" by overlap extension. Mutations can be introduced successfully by this method. Finally, the circular DNA is transformed into E. coli cells, where the nicks are ligated into a circular plasmid. One important requirement is that the parental plasmid carrying the target gene needs to be methylated by Dam methyltransferase or purified from Dam+ E. coli (i.e., DH5a).

Methods for In Vitro Mutagenesis.

Several different methodologies for mutagenesis have been developed to introduce mutations at predetermined sites or regions within mammalian genes. These methods of in vitro mutagenesis have had a transforming effect on the understanding of functions of protein, transcription regulatory elements, and noncoding RNAs and are now integral to molecular biology investigations. In this introduction, we summarize the most commonly used experimental approaches for mutagenesis and their major applications.

Random Mutagenesis Using Error-Prone DNA Polymerases.

"Random mutagenesis" is a technique that allows researchers to develop large libraries of variants of a particular DNA sequence. Once developed, these libraries can then be used for several purposes, including structure-function and directed evolution studies. Random mutagenesis is different from other mutagenesis techniques in that it does not require the researcher to have any prior knowledge about the structural properties of the DNA sequence being targeted, thus allowing for the unbiased discovery of novel or beneficial mutations. For this reason, random mutagenesis is especially useful for protein evolution studies. This protocol describes mutagenic replication in vitro by a low-fidelity DNA polymerase followed by selective polymerase chain reaction (PCR) amplification of the newly mutated sequences. The initial mutagenic DNA replication step is accomplished by heat-denaturing the template DNA and annealing primers possessing 5' extensions that are not complementary to the template. The purpose of the noncomplementary extensions on the primers is to allow for future selection of only the mutant strands. DNA replication is then performed by a low-fidelity DNA polymerase of choice (polymerase ß, η, or ι, or any combination of the three). After mutations have been incorporated into the template, the mutagenized strands are then selectively amplified using PCR. Selective amplification of the mutant strands is accomplished by performing a PCR procedure consisting of a first cycle with a low hybridization temperature followed by subsequent selection cycles under higher hybridization temperatures that do not allow amplification of the original unmutagenized template.

In Vitro Mutagenesis Using Double-Stranded DNA Templates: Selection of Mutants with DpnI.

In this protocol, two oligonucleotides are used to prime DNA synthesis by a high-fidelity polymerase on a denatured plasmid template. The two oligonucleotides both contain the desired mutation and have the same starting and ending positions on opposite strands of the plasmid DNA. The entire lengths of both strands of the plasmid DNA are amplified in a linear fashion during several rounds of thermal cycling, generating a mutated plasmid containing staggered nicks on opposite strands. Because of the amount of template DNA used in the amplification reaction, the background of transformed colonies containing wild-type plasmid DNA can be quite high unless steps are taken to enrich for mutant molecules. In this protocol, the products of the linear amplification reaction are treated with the restriction enzyme DpnI, which specifically cleaves fully methylated GMe6ATC sequences. DpnI will therefore digest the bacterially generated DNA used as template for amplification, but it will not digest DNA synthesized during the course of the reaction in vitro. DpnI-resistant molecules, which are rich in the desired mutants, are recovered by transforming E. coli cells to antibiotic resistance. Because the method works well with virtually any plasmid of moderate size (<7 kb), it can be used to introduce mutations directly into full-length cDNAs and eliminates the need for subcloning into specialized vectors.

Creating Insertions or Deletions Using Overlap Extension Polymerase Chain Reaction (PCR) Mutagenesis.

Overlap extension polymerase chain reaction (PCR) mutagenesis can be used for the generation of a specific point mutation, insertion, or deletion within a particular DNA sequence of interest. It requires relatively little preparation compared with other mutagenesis methods and does not require the use of restriction enzymes. Because of its versatility, the method has become widely used. Unlike methods of random mutagenesis, directed mutagenesis requires that the researcher already have a specific mutation in mind to implement. Traditional overlap extension PCR mutagenesis protocols remain limited in several critical ways, especially when it comes to generating insertions and deletions. For example, traditional protocols require that all sequence alterations be embedded within the primer itself, which makes it difficult to make insertions >30 nt. This protocol describes an overlap extension PCR mutagenesis method that is more versatile than its predecessors. Using this method, one can essentially make insertions and deletions of any size at any position within a given DNA sequence. To generate an insertion mutation, first prepare an insertion fragment and two flanking fragments by PCR. In the secondary PCR, the insertion fragment is recombined with two flanking fragments derived from the original template. This method can also be used to generate deletions, which is discussed in the latter part of the protocol.

Altered ß-Lactamase Selection Approach for Site-Directed Mutagenesis.

Many protocols exist to perform site-directed mutagenesis, and here we present one of the more commonly used ones-site-directed mutagenesis by altered ß-lactamase selection. ß-Lactamase is an enzyme that cleaves ampicillin, rendering it impotent to bacteria. Certain mutations in the active site of ß-lactamase can alter the substrate specificity of the enzyme and allow it to have increased hydrolytic activity for the cephalosporin family of antibiotics, a property not shared by wild-type lactamases. E. coli cells carrying the ß-lactamase triple mutant G238S:E240:R241G show increased resistance to cefotaxime and ceftriaxone, two cephalosporins, compared with wild-type cells. This protocol takes advantage of this property to select for plasmids that have undergone site-directed mutagenesis.

PEA15 regulates the DNA damage-induced cell cycle checkpoint and oncogene-directed transformation.

Regulation of the DNA damage response and cell cycle progression is critical for maintaining genome integrity. Here, we report that in response to DNA damage, COPS5 deubiquitinates and stabilizes PEA15 in an ATM kinase-dependent manner. PEA15 expression oscillates throughout the cell cycle, and the loss of PEA15 accelerates cell cycle progression by activating CDK6 expression via the c-JUN transcription factor. Cells lacking PEA15 exhibit a DNA damage-induced G2/M checkpoint defect due to increased CDC25C activity and, consequentially, higher cyclin-dependent kinase 1 (CDK1)/cyclin B activity, and accordingly they have an increased rate of spontaneous mutagenesis. We find that oncogenic RAS inhibits PEA15 expression and that ectopic PEA15 expression blocks RAS-mediated transformation, which can be partially rescued by ectopic expression of CDK6. Finally, we show that PEA15 expression is downregulated in colon, breast, and lung cancer samples. Collectively, our results demonstrate that tumor suppressor PEA15 is a regulator of genome integrity and is an integral component of the DNA damage response pathway that regulates cell cycle progression, the DNA-damage-induced G2/M checkpoint, and cellular transformation.