Effect of doubling volumes of PCR reagents

Effect of doubling volumes of PCR reagents

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We ran PCR with a positive control and two lanes of sample. We used the same sample, but in one PCR mixture we used a total volume of 50μL (25μL Taq, 5μL forward&reverse primer each, 10μL sample, 5μL dH2O) while in the other we halved each volume to a total of 25μL. Pipetting was done by the same person.

The positive control and the 25μL total volume lane gave nice and clear bands, but the 50μL sample showed nothing. How could this be explained?

Edit: We only ran one PCR with all the samples simultaneously, so no second thermocycler or different temperatures or anything. Didn't even consider that option!

Edit 2: In case it's relevant, we were using this:

The thermal cycler adjusts sample temperature based on the volume of the sample. So if you run samples of different volumes side by side, not all of them will cycle through the optimal temperatures of each step. If you were working with a "hot start" polymerase this is even more critical, as the Taq won't be able to amplify at all unless the sample is heated up to a specific temperature.

Reagent and laboratory contamination can critically impact sequence-based microbiome analyses

The study of microbial communities has been revolutionised in recent years by the widespread adoption of culture independent analytical techniques such as 16S rRNA gene sequencing and metagenomics. One potential confounder of these sequence-based approaches is the presence of contamination in DNA extraction kits and other laboratory reagents.


In this study we demonstrate that contaminating DNA is ubiquitous in commonly used DNA extraction kits and other laboratory reagents, varies greatly in composition between different kits and kit batches, and that this contamination critically impacts results obtained from samples containing a low microbial biomass. Contamination impacts both PCR-based 16S rRNA gene surveys and shotgun metagenomics. We provide an extensive list of potential contaminating genera, and guidelines on how to mitigate the effects of contamination.


These results suggest that caution should be advised when applying sequence-based techniques to the study of microbiota present in low biomass environments. Concurrent sequencing of negative control samples is strongly advised.


The Illumina sequencing platform [1], like other massively parallel sequencing platforms [2, 3], continues to produce ever-increasing amounts of data, yet suffers from under-representation and reduced quality at loci with extreme base compositions that are recalcitrant to the technology [1, 4–6]. Uneven coverage due to base composition necessitates sequencing to excessively high mean coverage for de novo genome assembly [7] and for sensitive polymorphism discovery [8, 9]. Although loci with extreme base composition constitute only a small fraction of the human genome, they include biologically and medically relevant re-sequencing targets. For example, 104 of the first 136 coding bases of the retinoblastoma tumor suppressor gene RB1 are G or C.

Traditional Sanger sequencing has long been known to suffer from problems related to the base composition of sequencing templates. GC-rich stretches led to compression artifacts. Polymerase slippage in poly(A) runs and AT dinucleotide repeats caused mixed sequencing ladders and poor read quality. Processes upstream of the actual sequencing, such as cloning, introduced bias against inverted repeats, extreme base-compositions or genes not tolerated by the bacterial cloning host. Gaps due to unclonable sequences had to be recovered and finished by PCR [10], or, in some cases, by resorting to alternative hosts [11]. Cloning bias hindered efforts to sequence the AT-rich genomes of Dictyostelium [12] and Plasmodium [13] and excluded the GC-rich first exons of about 10% of protein-coding genes in the dog (K Lindblad-Toh, personal communication) from an otherwise high-quality reference genome assembly [14].

New genome sequencing technologies [1–3, 15–17] no longer rely on cloning in a microbial host. Instead of ligating DNA fragments to cloning vectors, the three major platforms currently on the market (454, Illumina and SOLiD) involve ligation of DNA fragments to special adapters for clonal amplification in vitro rather than in vivo. Due to the massively parallel nature of the process, standardized reaction conditions must be applied to amplify and sequence complex libraries of fragments that comprise a wide spectrum of sequence compositions. All three platforms display systematic biases and unevenness as the observed coverage distributions are significantly wider than the Poisson distribution expected from unbiased, random sampling [18].

The Illumina sequencing process consists of i) library preparation on the lab bench, ii) cluster amplification, sequencing-by-synthesis and image analysis on proprietary instruments, followed by iii) post-sequencing data processing. Bias can be introduced at all three stages. For example, high cluster densities on the Illumina flowcell suppress GC-rich reads. Changes to sequencing kits, protocols and instrument firmware can affect the base composition of sequencing data. Moreover, bias is known to vary between laboratories, from run to run or even from lane to lane on the same flowcell. Such variability and instability in the system confound comparative studies [19, 20] and render systematic bias investigations difficult.

Here, we set out to evaluate sources of bias during Illumina library preparation and to ameliorate the effects. We undertook a systematic dissection of the process, using quantitative PCR (qPCR) instead of Illumina sequencing as a quick and system-independent read-out for base-composition bias. We identified library amplification by PCR as by far the most discriminatory step. We examined hidden factors such as make and model of thermocyclers and modified the thermocycling protocol. We tested alternative PCR enzymes and chemical ingredients in amplification reactions. Finally, we validated the qPCR results by Illumina sequencing. Our optimized protocol amplifies sequencing libraries more evenly than the standard protocol and minimizes the previously severe effects of PCR instrument and temperature ramp rate.

Betaine improves the co-amplification of the two alternatively spliced variants of the prostate-specific membrane antigen mRNA as well as the amplification of the coding cDNA region of c-jun. It is suggested that betaine improves the amplification of these genes by reducing the formation of secondary structure caused by GC-rich regions and, therefore, may be generally applicable to ameliorate the amplification of GC-rich DNA sequences.

Polymerase chain reaction (PCR) is a widely applied approach in molecular biology. GC-rich DNA sequences often require laborious work to optimize the amplification assay. Additives, DMSO ( 1 ) and glycerine ( 2 ), nucleotide analogs as 7-deaza dGTP ( 3 ) and dITP ( 4 ) or DNA template denaturation by NaOH ( 5 ) were introduced to optimize the amplification.

Co-amplifying both the prostate-specific membrane antigen (PSM) mRNA and the alternatively spliced variant (PSM′), differing by a deletion of 266 bases ( 6 ), by reverse transcription (RT) and PCR with one primer set located upstream (P1: 5′-AAACACTGCTGTGGTGGA) or downstream (P2: 5′-TAGCTCAACAGAATCCAGGC) from the deletion, we found that the ratio of PSM and PSM′ mRNA expressed in the prostatic cancer cell line LNCaP was <1 ( Fig. 1 ). This result is in contrast to data obtained by a RNase protection assay ( 6 ). Since the deletion contains GC-rich sequences (66% GC) the RT-PCR was optimized by both the involvement of 10% DMSO or 10% glycerine ( Fig. 1 ) and the application of PCR assays developed for fidelity and yield by selection of special Taq DNA polymerases. Unfortunately, these experimental approaches do not ameliorate the depressed amplification of PSM mRNA ( Fig. 2 ). However, the inclusion of betaine, purchased as monohydrate (Sigma), improves the amplification of PSM mRNA fundamentally ( Fig. 1 ). Betaine is effective with conventional Taq DNA polymerases (Promega Pharmacia Biotech) as well as with PCR assays designed for hot start PCR (AmpliTaq Gold™, Perkin Elmer) or for long and accurate (LA) PCR (Expand™ High Fidelity Taq, Boehringer Mannheim TaKaRa LA Taq, Boehringer Ingelheim Bioproducts) using, according to Barnes ( 7 ), a mixture of a minor proofreading enzyme and Taq DNA polymerase ( Fig. 2 ). Increasing betaine concentrations increase the signal intensity of the PSM amplicon and decrease that of the PSM′ amplicon. This relationship has similarities with the effect of increasing concentrations of an internal standard on the amplification efficiency of the target observed in the competitive PCR ( 8 ). Obviously, betaine makes PSM cDNA accessible for Taq DNA polymerase and favours, in this way, competition with the amplification of PSM′. The optimal betaine concentration for the amplification of the alternatively spliced PSM variants amounts to ∼1 M ( Fig. 3 ).

of additives on the amplification of alternatively spliced variants of PSM mRNA. cDNA synthesis with M-MLV reverse transcriptase (Promega), 1 ƒÊg RNA, isolated with TRIzol™Reagent (GIBCO BRL) and primer P2 at 50°C for 30 min. The amplification was performed with the 2.5 U Taq DNA polymerase (Promega) per 50 ml PCR assay, 1.7 mM MgCl 2 , 0.2 ƒÊM primer P1 and primer P2, hot-start and 25 cycles (annealing at 55°C, elongation at 72°C, denaturation at 95°C, each for 1 min) were used. PSM 511 bp, PSM′ 245 bp: lane 1, DNA marker XIV (Boehringer Mannheim) lane 2, without additives lane 3, 10% DMSO, 5 U Taq DNA polymerase were used since DMSO inhibits the enzyme ( 1 ) lane 4, 10% glycerine lane 5, 1 M betaine.

of additives on the amplification of alternatively spliced variants of PSM mRNA. cDNA synthesis with M-MLV reverse transcriptase (Promega), 1 ƒÊg RNA, isolated with TRIzol™Reagent (GIBCO BRL) and primer P2 at 50°C for 30 min. The amplification was performed with the 2.5 U Taq DNA polymerase (Promega) per 50 ml PCR assay, 1.7 mM MgCl 2 , 0.2 ƒÊM primer P1 and primer P2, hot-start and 25 cycles (annealing at 55°C, elongation at 72°C, denaturation at 95°C, each for 1 min) were used. PSM 511 bp, PSM′ 245 bp: lane 1, DNA marker XIV (Boehringer Mannheim) lane 2, without additives lane 3, 10% DMSO, 5 U Taq DNA polymerase were used since DMSO inhibits the enzyme ( 1 ) lane 4, 10% glycerine lane 5, 1 M betaine.

A further example of the effectiveness of betaine for the PCR of GC-rich sequences, is the improvement of the amplification of c-jun by a basic region containing 72% GC ( 1 ). For the amplification of the coding cDNA inserted in pBluescript ∼2.5 M betaine is optimal ( Fig. 3 ). Thus, the optimal betaine concentration seems to be dependent on the composition of the amplified DNA sequence.

Rees et al. ( 9 ) demonstrate that betaine reduces or even eliminates the base pair composition-dependent DNA thermal melting transition. Mytelka et al. ( 10 ), studying a novel class of DNA sequences that cause DNA polymerase to pause, observed that T7 DNA polymerase often recognizes pause sites near putative hairpin-loop structures. The addition of betaine relieves these pauses.

of PCR assays obtained from several suppliers by betaine. The PCR assays were performed as described by the manufacturers including the modifications outlined in Figure 1 and without or with 1 M betaine (lanes 3, 5, 7, 9, 11). Lane 1, DNA marker XIV (Boehringer Mannheim) lanes 2 and 3, Taq DNA polymerase (Promega) lanes 4 and 5, AmpliTaq Gold™ (Perkin Elmer) lanes 6 and 7, TaKaRa LA Taq (Boehringer Ingelheim Bioproducts) lanes 8 and 9, Taq DNA polymerase (Pharmacia Biotech) lanes 10 and 11, Expand™ High Fidelity Taq (Boehringer Mannheim).

of PCR assays obtained from several suppliers by betaine. The PCR assays were performed as described by the manufacturers including the modifications outlined in Figure 1 and without or with 1 M betaine (lanes 3, 5, 7, 9, 11). Lane 1, DNA marker XIV (Boehringer Mannheim) lanes 2 and 3, Taq DNA polymerase (Promega) lanes 4 and 5, AmpliTaq Gold™ (Perkin Elmer) lanes 6 and 7, TaKaRa LA Taq (Boehringer Ingelheim Bioproducts) lanes 8 and 9, Taq DNA polymerase (Pharmacia Biotech) lanes 10 and 11, Expand™ High Fidelity Taq (Boehringer Mannheim).

Effect of betaine on the amplification of both the alternatively spliced PSM variants and the c-jun cDNA fragment. PCR assay as described in Figure 1 including the following betaine concentrations: 0, 0.5, 1, 2.5 and 5 M (lanes 1–5 and 7–11). Lanes 1–5, PSM′ and PSM lanes 7–11, c-jun cDNA fragment inserted in pBluescript (50 ng), each 1 ƒÊM primer J1 (5′-ATGACTGCAAAGATGGAAACG) and primer J2 (5′-TCAAAATGAAAGCAACTGCTGCG), 30 cycles (annealing at 53°C, elongation at 72°C, denaturation at 95°C, each for 1 min) lane 6, DNA marker XIV (Boehringer Mannheim).

Effect of betaine on the amplification of both the alternatively spliced PSM variants and the c-jun cDNA fragment. PCR assay as described in Figure 1 including the following betaine concentrations: 0, 0.5, 1, 2.5 and 5 M (lanes 1–5 and 7–11). Lanes 1–5, PSM′ and PSM lanes 7–11, c-jun cDNA fragment inserted in pBluescript (50 ng), each 1 ƒÊM primer J1 (5′-ATGACTGCAAAGATGGAAACG) and primer J2 (5′-TCAAAATGAAAGCAACTGCTGCG), 30 cycles (annealing at 53°C, elongation at 72°C, denaturation at 95°C, each for 1 min) lane 6, DNA marker XIV (Boehringer Mannheim).

In summary, the application of betaine in the PCR assay improves the amplification of both the PSM and c-jun and may be generally applicable to ameliorate the amplification of GC-rich DNA sequences.

While this paper was under review, we became aware of an independent study of Baskaran et al. ( 11 ) in which betaine is applied in combination with DMSO for the uniform amplification of a mixture of DNA with varying GC content. Hengen ( 12 ), referring to an information sheet for LA-PCR provided by Wayne Barnes, pointed out that 1.3 M betaine and 1.3% DMSO added to LA-PCR mixtures improves processivity.

The PCR method is extremely sensitive, requiring only a few DNA molecules in a single reaction for amplification across several orders of magnitude. Therefore, adequate measures to avoid contamination from any DNA present in the lab environment (bacteria, viruses, or human sources) are required. Because products from previous PCR amplifications are a common source of contamination, many molecular biology labs have implemented procedures that involve dividing the lab into separate areas. [1] One lab area is dedicated to preparation and handling of pre-PCR reagents and the setup of the PCR reaction, and another area to post-PCR processing, such as gel electrophoresis or PCR product purification. For the setup of PCR reactions, many standard operating procedures involve using pipettes with filter tips and wearing fresh laboratory gloves, and in some cases a laminar flow cabinet with UV lamp as a work station (to destroy any extraneomultimer formation). PCR is routinely assessed against a negative control reaction that is set up identically to the experimental PCR, but without template DNA, and performed alongside the experimental PCR.

Secondary structures in the DNA can result in folding or knotting of DNA template or primers, leading to decreased product yield or failure of the reaction. Hairpins, which consist of internal folds caused by base-pairing between nucleotides is inverted repeats within single-stranded DNA, are common secondary structures and may result in failed PCRs.

Typically, primer design that includes a check for potential secondary structures in the primers, or addition of DMSO or glycerol to the PCR to minimize secondary structures in the DNA template, [2] are used in the optimization of PCRs that have a history of failure due to suspected DNA hairpins.

Taq polymerase lacks a 3′ to 5′ exonuclease activity. Thus, Taq has no error-proof-reading activity, which consists of excision of any newly misincorporated nucleotide base from the nascent (i.e., extending) DNA strand that does not match with its opposite base in the complementary DNA strand. The lack in 3′ to 5′ proofreading of the Taq enzyme results in a high error rate (mutations per nucleotide per cycle) of approximately 1 in 10,000 bases, which affects the fidelity of the PCR, especially if errors occur early in the PCR with low amounts of starting material, causing accumulation of a large proportion of amplified DNA with incorrect sequence in the final product. [3]

Several "high-fidelity" DNA polymerases, having engineered 3′ to 5′ exonuclease activity, have become available that permit more accurate amplification for use in PCRs for sequencing or cloning of products. Examples of polymerases with 3′ to 5′ exonuclease activity include: KOD DNA polymerase, a recombinant form of Thermococcus kodakaraensis KOD1 Vent, which is extracted from Thermococcus litoralis Pfu DNA polymerase, which is extracted from Pyrococcus furiosus and Pwo, which is extracted from Pyrococcus woesii. [4]

Magnesium is required as a co-factor for thermostable DNA polymerase. Taq polymerase is a magnesium-dependent enzyme and determining the optimum concentration to use is critical to the success of the PCR reaction. [5] Some of the components of the reaction mixture such as template concentration, dNTPs and the presence of chelating agents (EDTA) or proteins can reduce the amount of free magnesium present thus reducing the activity of the enzyme. [6] Primers which bind to incorrect template sites are stabilized in the presence of excessive magnesium concentrations and so results in decreased specificity of the reaction. Excessive magnesium concentrations also stabilize double stranded DNA and prevent complete denaturation of the DNA during PCR reducing the product yield. [5] [6] Inadequate thawing of MgCl2 may result in the formation of concentration gradients within the magnesium chloride solution supplied with the DNA polymerase and also contributes to many failed experiments . [6]

PCR works readily with a DNA template of up to two to three thousand base pairs in length. However, above this size, product yields often decrease, as with increasing length stochastic effects such as premature termination by the polymerase begin to affect the efficiency of the PCR. It is possible to amplify larger pieces of up to 50,000 base pairs with a slower heating cycle and special polymerases. These are polymerases fused to a processivity-enhancing DNA-binding protein, enhancing adherence of the polymerase to the DNA. [7] [8]

Other valuable properties of the chimeric polymerases TopoTaq and PfuC2 include enhanced thermostability, specificity and resistance to contaminants and inhibitors. [9] [10] They were engineered using the unique helix-hairpin-helix (HhH) DNA binding domains of topoisomerase V [11] from hyperthermophile Methanopyrus kandleri. Chimeric polymerases overcome many limitations of native enzymes and are used in direct PCR amplification from cell cultures and even food samples, thus by-passing laborious DNA isolation steps. A robust strand-displacement activity of the hybrid TopoTaq polymerase helps solve PCR problems that can be caused by hairpins and G-loaded double helices. Helices with a high G-C content possess a higher melting temperature, often impairing PCR, depending on the conditions. [12]

Non-specific binding of primers frequently occurs and may occur for several reasons. These include repeat sequences in the DNA template, non-specific binding between primer and template, high or low G-C content in the template, or incomplete primer binding, leaving the 5' end of the primer unattached to the template. Non-specific binding of degenerate primers is also common. Manipulation of annealing temperature and magnesium ion concentration may be used to increase specificity. For example, lower concentrations of magnesium or other cations may prevent non-specific primer interactions, thus enabling successful PCR. A "hot-start" polymerase enzyme whose activity is blocked unless it is heated to high temperature (e.g., 90–98˚C) during the denaturation step of the first cycle, is commonly used to prevent non-specific priming during reaction preparation at lower temperatures. Chemically mediated hot-start PCRs require higher temperatures and longer incubation times for polymerase activation, compared with antibody or aptamer-based hot-start PCRs. [ citation needed ]

Other methods to increase specificity include Nested PCR and Touchdown PCR.

Computer simulations of theoretical PCR results (Electronic PCR) may be performed to assist in primer design. [13]

Touchdown polymerase chain reaction or touchdown style polymerase chain reaction is a method of polymerase chain reaction by which primers will avoid amplifying nonspecific sequences. The annealing temperature during a polymerase chain reaction determines the specificity of primer annealing. The melting point of the primer sets the upper limit on annealing temperature. At temperatures just below this point, only very specific base pairing between the primer and the template will occur. At lower temperatures, the primers bind less specifically. Nonspecific primer binding obscures polymerase chain reaction results, as the nonspecific sequences to which primers anneal in early steps of amplification will "swamp out" any specific sequences because of the exponential nature of polymerase amplification.

The earliest steps of a touchdown polymerase chain reaction cycle have high annealing temperatures. The annealing temperature is decreased in increments for every subsequent set of cycles (the number of individual cycles and increments of temperature decrease is chosen by the experimenter). The primer will anneal at the highest temperature which is least-permissive of nonspecific binding that it is able to tolerate. Thus, the first sequence amplified is the one between the regions of greatest primer specificity it is most likely that this is the sequence of interest. These fragments will be further amplified during subsequent rounds at lower temperatures, and will out compete the nonspecific sequences to which the primers may bind at those lower temperatures. If the primer initially (during the higher-temperature phases) binds to the sequence of interest, subsequent rounds of polymerase chain reaction can be performed upon the product to further amplify those fragments.

Annealing of the 3' end of one primer to itself or the second primer may cause primer extension, resulting in the formation of so-called primer dimers, visible as low-molecular-weight bands on PCR gels. [14] Primer dimer formation often competes with formation of the DNA fragment of interest, and may be avoided using primers that are designed such that they lack complementarity—especially at the 3' ends—to itself or the other primer used in the reaction. If primer design is constrained by other factors and if primer-dimers do occur, methods to limit their formation may include optimisation of the MgCl2 concentration or increasing the annealing temperature in the PCR. [14]

Deoxynucleotides (dNTPs) may bind Mg 2+ ions and thus affect the concentration of free magnesium ions in the reaction. In addition, excessive amounts of dNTPs can increase the error rate of DNA polymerase and even inhibit the reaction. [5] [6] An imbalance in the proportion of the four dNTPs can result in misincorporation into the newly formed DNA strand and contribute to a decrease in the fidelity of DNA polymerase. [15]


Anticipated Results—

Females possess only X chromosomes, and therefore a single band of 977 bp is anticipated, corresponding to the expected size of the PCR product from the X chromosome (Fig. 2, lane 1). Males possess both X and Y chromosomes, and therefore amplification by PCR will produce two DNA bands of 977 bp and 788 bp (Fig. 2, lane 2). The control DNA provided with the kit produces either one band, indicating a female DNA donor, or two bands, indicating a male (Fig. 2, lane 3). Typically, the DNA bands representing PCR products from the control DNA are more intense than the products from the student-isolated DNA, presumably because there is less DNA template in the student samples. Faint bands greater than 1000 bp in size occasionally appear due to nonspecific binding of the primers.

The control DNA is used to verify that the PCR reagents are functional, but can also serve as an unknown. The results for an additional DNA donor (such as the teaching assistant or instructor) can be provided to the students as a second unknown (Fig. 2, lane 5). Students can determine the gender of unknown DNA donors by comparison to known samples or by the sizes of the PCR products relative to the DNA standards. Students should estimate the size of the X and Y chromosome fragments using semi-logarithmic paper. Plotting the size of the DNA standards on the logarithmic axis versus the Rf value (or distance migrated on the gel) on the x-axis typically results in a straight line.

Common Student Pitfalls—

In each of the four semesters that the exercise has been offered, some of the groups have obtained the anticipated results. Among the groups demonstrating only partial success, many obtained visible bands in the lanes containing control DNA and molecular size standards but lacked bands in one (or both) of the student-derived samples. This can result from starting with a very small sample of cheek cells or from errors in isolating DNA (such as discarding the BD-4 supernatant that contains the DNA). Another student pitfall involves inaccurate pipetting technique, particularly while drawing up liquid solutions. Because PCR is highly sensitive to the concentrations of its reaction components, pipetting errors can lead to the appearance of extra bands on the gel or to complete failure of the PCR, even with the control DNA. A common warning sign of pipetting inaccuracy is that a group may run out of their solution aliquots during preparation of the DNA samples.

To maximize the probability of student success, it is recommended that the exercise be prerun with all of the actual kits, enzymes, and solutions to be used by the students. Prerunning the exercise will also enable the teaching assistant to optimize the volumes of PCR product loaded onto the gel as well as the time period for running the gels.

Presentation of Data—

We have used two different formats for student presentation of data. The first is a worksheet or short report in which students respond to the following instructions. First, attach a copy of the PCR data to the report second, generate a standard curve on semi-logarithmic paper and estimate the sizes of the PCR products third, discuss the results using the following questions as a guide. Did you obtain PCR products from each of your samples? How many bands did you get from amplification of the control DNA? What size fragments did you obtain from the amplification of your samples?

The second format is a short formal laboratory report (two to three typed, double-spaced pages) requiring a title, statement of objectives, materials and methods (referencing the manual with technical changes noted), results (including labeled figures with titles and legends), discussion, and references. In the discussion, students should explain the banding pattern of known samples, identify the gender of the unknown DNA donor, discuss whether the results were consistent with anticipated results, propose explanations for discrepancies (using theoretical knowledge of trouble-shooting strategies), and describe how the exercise might be improved in the future. An interesting observation in one of our classes has been that many of the students are puzzled by the appearance of two bands in the male sample. The answer can be drawn out of students by asking them to explain the chromosomal composition of males (XY) versus females (XX).

Effects of OTU Clustering and PCR Artifacts on Microbial Diversity Estimates

Next-generation sequencing has increased the coverage of microbial diversity surveys by orders of magnitude, but differentiating artifacts from rare environmental sequences remains a challenge. Clustering 16S rRNA sequences into operational taxonomic units (OTUs) organizes sequence data into groups of 97 % identity, helping to reduce data volumes and avoid analyzing sequencing artifacts by grouping them with real sequences. Here, we analyze sequence abundance distributions across environmental samples and show that 16S rRNA sequences of >99 % identity can represent functionally distinct microorganisms, rendering OTU clustering problematic when the goal is an accurate analysis of organism distribution. Strict postsequencing quality control (QC) filters eliminated the most prevalent artifacts without clustering. Further experiments proved that DNA polymerase errors in polymerase chain reaction (PCR) generate a significant number of substitution errors, most of which pass QC filters. Based on our findings, we recommend minimizing the number of PCR cycles in DNA library preparation and applying strict postsequencing QC filters to reduce the most prevalent artifacts while maintaining a high level of accuracy in diversity estimates. We further recommend correlating rare and abundant sequences across environmental samples, rather than clustering into OTUs, to identify remaining sequence artifacts without losing the resolution afforded by high-throughput sequencing.

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  • 1. Choosing a knockout strategy
  • 2. Selecting gRNA target sites and performing vector cloning
  • 3. Introducing gRNAs by transfection or transduction
  • 4. Isolation and expansion of single-cell clones
  • 5. Knockout verification by western blot analysis, PCR, and/or Sanger sequencing.

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The superfamily of nuclear receptors include ligand-activated transcription factors, which can respond to hormonal or metabolic ligands. Because these proteins regulate the transcription of target genes and thereby change mRNA levels, their activities are particularly amenable to study by the methodology of real-time RT-PCR.

Peroxisome proliferator-activated receptor (PPAR)-α, -β (-δ), and -γ are members of the nuclear receptor superfamily and known to be activated by endogenous saturated and unsaturated long-chain fatty acids, eicosanoids, and prostaglandins. In addition, two classes of antidiabetic agents are known to bind PPARs: fibrates (e.g., fenofibrate, known clinically as Lofibra or Tricor) bind PPAR-α, whereas thiazolidinediones (e.g., rosiglitazone, known clinically as Avandia) bind PPAR-γ. The binding of these drugs activates the respective transcription factor to enhance the transcription of target genes and effect a physiological response.

Because PPAR-α activation favors fatty acid oxidation in tissues (especially the liver and heart), it may enhance insulin sensitivity by reducing intracellular fatty acid accumulation. PPAR-γ activation favors the storage of lipids in adipose tissue, thereby protecting the rest of the body from lipid overload and insulin resistance (15). Activation of PPARs may also enhance insulin sensitivity in other ways. For example, PPAR-γ is known to upregulate adiponectin (5, 52). Adiponectin is considered to be an “adipocytokine” because it is made exclusively by adipose tissue and then secreted into the circulation. Plasma adiponectin levels appear to be inversely correlated with obesity and insulin resistance. Adiponectin action on tissues became better understood when two receptors for adiponectin were recently identified (51). Adiponectin receptor 1 (AdipoR1) is expressed ubiquitously but most highly in skeletal muscle, whereas AdipoR2 is primarily expressed in the liver (51). Because the adiponectin system is already implicated in enhancing insulin sensitivity, we wanted to know whether insulin sensitization by PPARs could in part be due to their potential effects on adiponectin receptor expression.

In these experiments, we sought to determine the impact of various nuclear receptor agonists (drugs) and a gene knockout (genes) on the expression of adiponectin receptors in the liver (Fig. 6). First, mice were fed standard chow supplemented with 30 mg/kg LG268 [retinoid X receptor (RXR)], 0.5% (wt/wt) fenofibrate (PPAR-α), 150 mg/kg troglitazone (PPAR-γ), 0.05% (wt/wt) prenenolone-16α-carbonitrile [pregnane X receptor (PXR)], 3 mg/kg TCPOBOP (constitutive androstone receptor), 0.5% (wt/wt) chenodeoxycholic acid (farnesoid X receptor), 50 mg/kg T1317 [liver X receptor (LXR)], or 30 mg/kg LG268 + 50 mg/kg T1317 (RXR+LXR) for 12 h. Second, both wild-type and PPAR-α knockout mice were fed a standard chow with or without 0.5% (wt/wt) fenofibrate for 7 days. Animal experiments were approved by the Institutional Animal Care and Use Committee of the University of Texas Southwestern Medical Center (Dallas, TX). Tissues were harvested, RNA was extracted, real-time PCR was performed to determine the relative abundance of mRNA (25), and calculations were done using the comparative CT method (User Bulletin No. 2, Perkin-Elmer Life Sciences). Data were procured by analyzing the expression in individual livers (triplicate measurement) in each group (n = 3–4) and are expressed as means ± SE. Thus the SE reflects biological variation in addition to measurement variation. In the first experiment, AdipoR1 mRNA levels were mildly increased by activation of RXR and PPAR-α (1.34 ± 0.15- and 1.70 ± 0.12-fold), as were AdipoR2 mRNA levels (1.70 ± 0.07- and 2.31 ± 0.24-fold). Notably, the PPAR-γ agonist had no effect. In the second experiment, AdipoR1 and AdipoR2 mRNA expression were increased 1.91 ± 0.05- and 3.28 ± 0.10-fold, respectively, in wild-type animals treated with fenofibrate. No increase was observed for either gene in treated PPAR-α knockout animals. Similar results were observed in livers of mice treated with another PPAR-α agonist, GW7647, by oral gavage [2 doses of 5 mg/kg over 14 h data not shown (7)].

Fig. 6.Example of real-time PCR analysis. Mice received drugs that specifically activate various nuclear hormone receptors (listed on the x-axis). Liver RNA was prepared, and the RNA levels for adiponectin receptors 1 and 2 (AdipoR1 and AdipoR2) were measured using an ABI Prism 7900HT system, the SYBR green I method, and target-specific primers (AdipoR1: forward 5′-cagtgggaccggtttgc-3′ and reverse 5′-aagccaagtcccaggaacac-3′ AdipoR2: forward 5′-aacgaatggaagagtttgtttgtaa-3′ and reverse 5′-gtagcacatcgtgagggatca-3′). Full details of the method can be found in Ref. 25, and cyclophilin was used as the invariate housekeeping gene. Left: only peroxisome proliferator-activated receptor (PPAR)-α (solid bars) and its dimeric partner retinoid X receptor (RXR hatched bars) elicit an increase in hepatic AdipoR1 and AdipoR2 mRNA levels compared with the control group (*P < 0.05 and ***P < 0.001). Right: the importance of PPAR-α is highlighted, because in mice lacking this nuclear hormone receptor (Ppar-α −/− ) the fibrate drug fails to work. a,b By ANOVA, groups sharing a common letter designation are not different. PXR, pregnane X receptor CAR, constitutive androstane receptor FXR, farnesoid X receptor LXR, liver X receptor.

These results demonstrate that fenofibrate enhances adiponectin receptor expression in the liver by a PPAR-α-dependent mechanism and suggest that fenofibrate may enhance insulin sensitivity by increasing adiponectin action on the liver. Further investigation is needed to determine whether PPAR-α is acting directly on the promoters of AdipoR1 and AdipoR2 genes in liver cells or indirectly by some other means (e.g., changes in metabolism). Also, this result is consistent with the therapeutic potential of PPAR-α/γ dual agonists (5, 52).

This example is a testimony to the power of real-time PCR in elucidating molecular events that underlie physiology. By applying a single methodology, that of real-time PCR, to a drug-treated mouse knockout model, we were able to gain insight into novel molecular mechanisms potentially involved in insulin sensitization. Obviously, the ability to precisely detect relative changes in gene expression is a valuable tool for studying any number of physiological, pathophysiological, and developmental models. Thus real-time PCR can be a powerful first step in many biomedical research projects and programs.

Watch the video: PCR -1 Lecture - Professor B. Albrecht (May 2022).