Chai FAQs

Real-time PCR reactions are the default setting for the Open qPCR system, but the instrument is also capable of functioning as a conventional/endpoint thermocycler by making a few adjustments in the protocol editor while setting up an experiment.

How to use Open qPCR for conventional PCR:

After creating a new experiment, the default protocol appears and the default real-time setting is indicated by the amplification icon (green arrow):

To switch to endpoint reactions, turn off Gather Data (GATHER DATA) for all steps & ramps in all stages of the protocol.

Toggle the STEP & RAMP options from the green ON position:

to the gray OFF position:

Confirm that the setting has been changed from real-time to endpoint by verifying that there are no amplification icons visible for any of the steps in the protocol.

Consequences of switching to conventional PCR

Choosing to turn off the data gathering functions will cause a few other changes that are important to keep in mind when deciding if switching to endpoint reactions is the right option. Using real-time settings, the results screen displays a generated amplification curve and Cq values if a dye or probe is used in the reaction.

Using conventional PCR reactions with no fluorescence detection, the LEDs will not be activated throughout the duration of the protocol. As a result, no amplification curve or Cq values will be generated (if using a dye or probe) and these areas will be blank on the results screen.

The Superattenuator Yeasts Test Kit detects both Diastaticus and Brettanomyces. There are two options for enrichment that differ in terms of enrichment time and sensitivity, as Brettanomyces typically requires a longer incubation time than Diastaticus to achieve single cell detection when followed by qPCR.

To reach a limit of detection of 1 cell/sample when followed by qPCR for Diastaticus and greater sensitivity than direct qPCR for Brettanomyces, perform a single 48 hour sample enrichment using FastOrange Wild Yeast solution and proceed with analysis via qPCR.

When single cell detection for both Diastaticus and Brettanomyces is required, enrich samples separately for 72 hours in FastOrange Brett Broth for Brettanomyces and 48 hours in FastOrange Wild Yeast Broth for Diastaticus. Then combine both enriched samples into a single 1.5 mL tube and proceed with analysis by qPCR.

Enriching samples for Diastaticus requires a 48-hour enrichment using the FastOrange Wild Yeast solution, which provides a limit of detection of 1 cell/sample in 48 hours when followed by qPCR. A volume of 40 mL is recommended for sensitive detection. Less volume of FastOrange Wild Yeast solution may be used, but this will result in decreased sensitivity. Diastaticus grows in aerobic environments, so lay your enrichment containers on the side to maximize air exposure when possible. All enrichment solution and sample volume should be transferred utilizing sterile technique.

Using the 40 mL FastOrange Wild Yeast Enrichment Bottles:

Add 40 mL of your sample to a 40 mL FastOrange Wild Yeast Enrichment Bottle and incubate for 48 hours at room temperature (25 C +/- 2 C). Lay the bottle flat on its side and vent the sample to avoid pressure buildup.

Using the 240 mL FastOrange Wild Yeast Broth:

Aliquot your entire 240 mL FastOrange Wild Yeast Broth from the glass bottle into individual enrichment containers with your chosen volume. We recommend 40 mL for sensitive detection. Use sterile technique to transfer the solution. Aliquoted enrichment containers can be stored at room temperature (max 25 C), protected from light.

Add your sample in a 1:1 ratio to one enrichment container, again using sterile technique. Incubate the enrichment container for 48 hours at room temperature (25 C +/- 2 C). Leave the container on its side and vent the sample to avoid pressure buildup.

After your sample has been enriched for 48 hours, proceed with analysis by qPCR.

The 240 mL FastOrange Broth bottles can be used for sample enrichment as long as quantities are scaled appropriately and solution is transferred using sterile technique to prevent contamination. It is recommended to aliquot the entire 240 mL broth bottle into individual enrichment containers at one time to minimize contamination. Unused enrichment containers can be stored at room temperature (max 25 C) protected from light.

Determine what enrichment containers to use. We recommend the following types of sterile enrichment containers for FastOrange enrichment:

T-75 cell culture flasks: Recommended for most enrichments as one-use containers. We suggest using these for standard 50 mL volume enrichments.

Reusable glass bottles: Recommended as a cost-effective alternative if an autoclave is available

50 mL Falcon/Eppendorf centrifuge tubes: Recommended for enriching bacteria growing in anaerobic environments such as Megasphaera, Pectinatus and L. acetotolerans* or general purpose 25 mL volume enrichments.

15 mL tubes: Recommended for low-cost enrichments where the highest degree of sensitivity is not required.

In general, use containers that will stand vertically to minimize air exposure when enriching using FastOrange B or FastOrange Brett Broth. When enriching using FastOrange Yeast or Wild Yeast Broth, find a container that can be laid flat on its side for maximum air exposure.

Aliquot the FastOrange Broth into your enrichment containers. Carefully aliquot the desired quantity of solution as necessary until you have transferred your total required volume of broth (typically 40 50 mL) into each enrichment container based on graduations in the receiving container. Store the containers vertically at room temperature (max 25 C), protected from light.

Add a volume of sample in a 1:1 ratio with the FastOrange Broth in one enrichment container using sterile technique. Transfer the total required volume of sample into the enrichment container (i.e., if you have a cell culture flask with 40 mL enrichment of FastOrange enrichment solution, pour in 40 mL of sample). Incubate according to the instructions of the specific enrichment broth.

General Notes:

Store FastOrange Broth in the dark at room temperature (max 25 C). If youre using the broth to detect beer spoilers via color change only, protect from light during the sample enrichment. If youre following up the enrichment with analysis via PCR, the enrichment container does not have to be protected from light during sample enrichment.

*For Megasphaera, Pectinatus and L. acetotolerans, please refer to the article on how to enrich for anerobic bacteria.

Understanding the differences between the single channel and dual channel open qPCR instruments will allow you to decide which instrument is the best option for your needs. The number of target sequences your assay design needs to amplify and the fluorophores required by your test kit are important to consider, along with the desired level of monetary investment.

Single Channel Open qPCR

One channel

One target sequence detection/reaction tube

16 wells

Wavelength detection range: 513 nm 555 nm

Single channel open qPCR instruments are a great choice when minimizing cost is your top concern. However, it is important to keep in mind that a single channel instrument will significantly limit the capabilities of your assay, as detection for only one target sequence may be performed in each reaction tube.

Dual Channel Open qPCR

Two channels

Two target sequence detection/reaction tube

16 wells

Wavelength detection ranges: 508 nm 532 nm & 573 nm 597 nm

Dual channel open qPCR instruments are ideal for any operation wishing for multiplexing capabilities, which offers many advantages. The two channels in these instruments allow for detection of two target sequences in each reaction tube. If your assay requires amplification of multiple target sequences, using multiplex PCR rather than several singleplex PCR reactions is much more efficient, as it maximizes the use of limited starting material and lowers reagent costs. Instruments with multiplexing capabilities are ideal for high-volume environments that need to perform large quantities of the same test repeatedly. Additionally, a multiplexed channel may be used as an internal control for your assay to confirm the absence of reaction inhibitors or verify the completion of the extraction process.

Model

Compatible Fluorophores

Single Channel

FAM

SYBR Green

Chai Green

Dual Channel

FAM

SYBR Green

Chai Green

HEX

VIC

JOE

A PCR master mix is a premixed solution that contains most of the components necessary to run a PCR assay. The mix contains Taq DNA polymerase, dNTPs, MgCl2, as well as enhancers and stabilizers in a buffer that is optimized for DNA amplification by PCR.

Advantages of Using a PCR Master Mix

The major appeal for using a PCR master mix is its convenience. Most of the components have already been added and the formula has already been optimized, therefore significantly reducing pre-assay preparation time.

The use of master PCR mixes also ensures a high degree of consistency, even in high-volume assay environments, and the fewer pipetting steps involved also means fewer opportunities for contamination. Using a PCR master mix also reduces the chance for a preparation error, such as accidentally leaving out a component.

Additionally, commercial master mixes may contain various enhancers and stabilizers not familiar to novice PCR users. They are also subject to quality control procedures and analysis that produce an extremely accurate and reliable product. These factors result in overall enhanced assay performance.

All of these advantages are especially important to consider for users new to PCR.

What Do I Add to a PCR Master Mix?

Primers

Detection Dye or Probes

Sample

RNAse-free water

Master Mix Format & Stability

PCR master mixes commonly exist in two major formats. Most often, PCR master mixes come in liquid format. Liquid PCR master mixes generally require storage conditions between -20 C 4 C, and tend to be less expensive than lyophilized mixes. They are thawed before use in a PCR assay.

PCR master mixes may also come in a lyophilized, or freeze-dried format. This allows the mix to be shipped at ambient temperatures. Some lyophilized master mixes may also be stored at ambient temperatures long-term. The master mix is then reconstituted with the accompanying buffer solution before use.

Routine PCR Master Mixes

Master mixes used for routine PCR assays typically amplify target sequences up to 5 kb in size, with a GC content ranging between 40% and 60%.

Various master mixes also exist for assays desiring an enhanced level of PCR performance:

Hot Start Taq: prevention of non-specific amplification at lower temperatures

High-Fidelity Taq: specialized enzymes for long-range amplicons, up to 20 kb

Real-Time PCR Master Mixes

Master mixes created for use with Real-time PCR assays are optimized for assays employing either dye or probe detection chemistry.

PCR master mixes used in assays employing dye detection chemistry may or may not contain the actual double-stranded DNA binding dye. If the dye is not included, the user may add an external custom dye to the master mix.

Similarly, PCR master mixes designed for assays utilizing probe detection chemistry do not include the fluorophore-labeled probe in the mix, but it may be added by the user.

PCR master mixes intended for assays using either type of detection chemistry may also include a passive reference dye; the most commonly included is ROX. This reference dye is necessary for the operation of certain, but not all real-time PCR instruments. Whether your instrument requires this reference dye should be determined prior to choosing a real-time PCR master mix.

Should I Ever Use a Home-Brewed PCR Master Mix?

In most cases, the advantages of using a PCR master mix are far greater than the benefits that may be gained from creating a personalized PCR master mix solution.

However, for assays requiring an enhanced level of user control over the experimental conditions, home-brewed PCR master mixes may be the superior choice. Downstream applications demanding this level of user control over the assay specifications may include sequencing, cloning, etc.

These home-brewed PCR master mixes often come at a reduced cost as well. Still, buffer optimization must be performed by the user, which involves an additional time investment.

PCR (Polymerase Chain Reaction) is a method to selectively copy ("amplify") a segment of DNA. By repeatedly duplicating the DNA, PCR can generate billions of copies from even a single starting molecule. Because only the desired segment of DNA is amplified, PCR isolates a "needle in a haystack", allowing minute quantities of pathogens to be detected and providing abundant DNA for other molecular biology techniques.

In PCR, a sample undergoes a series of heating and cooling cycles in the presence of DNA polymerase, an enzyme that replicates DNA. During every cycle, each existing copy of target DNA serves as a template for synthesizing a new copy. This chain reaction doubles the number of target DNA each cycle; an exponential amplification. PCR reactions typically involve 30 - 40 cycles which are automated by a PCR thermocycler.

PCR is widely used to diagnose infectious diseases and hereditary disorders. It is also used in veterinary diagnostics, food safety testing, water & beverage quality monitoring, and forensics. In research, PCR is used to monitor gene expression, perform gene cloning & manipulation, and prepare DNA for sequencing.

Singleplex PCR uses one pair of primers to amplify a single target. Multiplex PCR uses several pairs of primers to amplify more than one target sequence. Both are performed in a single reaction tube. Multiplex PCR is often the preferred option due to decreased reaction time and efficient use of both starting material and reagents.

PCR Multiplexing Methods

Optical Channel Multiplexing

The most common way of performing PCR multiplexing is through the use of various optical channels. Separate probes specific to each target sequence are each labeled with a unique combination of fluorophores. The use of these unique fluorophores allows different target sequences to be detected in separate optical channels.

The number of optical channels needed depends on the number of different target sequences to be detected within the reaction. Instruments supporting 2-4 optical channels are commonly available.

Optical channel multiplexing is only compatible with an assay utilizing probe detection chemistry. It is not suitable with a dye-based detection assay, due to the non-specific product amplification by the DNA binding dye. The dye intercalates with all double-stranded DNA, making distinction between the fluorescent signals of different target sequences impossible.

Screening Assays

Screening assays use multiple target-specific primers, but detect the fluorescence emitted from amplicons through the use of only one optical channel. This means that all of the target sequences, if present in the sample, will produce fluorescent signals that will be detected within that same single optical channel.

The results do not distinguish between different amplified products. However, screening assays save time and reduce costs since only the samples that have been flagged as positive for a target sequence will undergo further analysis. These assays are compatible with either intercalating dye or probe detection chemistries.

Screening assays are typically used in situations where there are many samples to be tested, each for the presence of multiple targets, such as a food/beverage quality control assay that screens for multiple pathogens.

TmMultiplexing

While melt curve analysis (MCA) is typically used to detect non-specific product amplification, it may also be used for multiplexing.

MCA is applied to the identification of several specific PCR products by measuring the decrease in fluorescence intensity as the double-stranded DNA dissociates into single-stranded DNA. The temperature at which 50% of the total double-stranded DNA present separates is referred to as melting temperature (Tm). The assay is designed such that every amplicon has a different melting temperature. Tmmultiplexing may then be achieved by differentiating PCR products using the varying melting temperatures of the different amplicons.

Using Tmmultiplexing, the MCA data below shows the differentiation of GAPDH and HPRT amplicons, with each peak corresponding to a specific amplicon.

Melt Curve Analysis of HPRT & GAPDH Reaction Products via Eva Green Assay

Singleplex PCR Advantages + Disadvantages

+ Ease of Design

With a single target assay, there is no competition for reaction components, making assay design much easier.

+ Decreased Instrument Cost

Only a single channel real-time PCR instrument is required for singleplex assays.

+ Choice of Detection Chemistry

Singleplex assays may be performed using either a dye or probe detection chemistry, affording the user additional control over assay specifications.

- Inefficient for Multiple Target Assays

Using several singleplex reactions to perform a multiple target sequence assay means an increase in reagent quantity consumption and sample material requirements. Additional costs and labor are incurred as a result.

Multiplex PCR Advantages + Disadvantages

+ Maximum Efficiency

Using multiplex PCR rather than several singleplex PCR reactions maximizes the use of limited starting material and lowers reagent costs for assays requiring amplification of several target sequences.

+ Internal Control Option

An internal control coupled with the sample confirms the absence of inhibitors in the reaction. Internal controls may also serve to verify that the extraction process was complete.

+ SNP Genotyping via Allelic Discrimination Plot

A multiplexed end-point assay for detecting variants of a single nucleotide sequence would classify unknown samples as either homozygous or heterozygous and would show up as clusters in the allelic discrimination plot.

+ Useful for High-Volume Environments

Multiplex assays are excellent for labs needing to perform a large quantity of the same test repeatedly.

- Cross-Reactivity

A non-optimized multiplex assay may experience primer-primer hybridization or incorrect primer-template binding. This produces inaccurate Cq(quantification cycle) values.

- Multiple Optimization Rounds

Competition for reaction components causes low-abundance targets to be outcompeted by high-abundance targets. Multiple rounds of optimization are often needed to set the appropriate primer concentration.

Yes, the PCR strip tubes and strip caps may be cut to any length and used in the Open qPCR instrument. Ensure that the cutting device used is sterilized with 70% ethanol to prevent any carryover contamination. Adhere to Good Laboratory Practices (GLP).

Two major types of chemistries may be utilized to detect the products of your qPCR cycles, each with particular advantages. Understanding the differences between the two will ensure you choose the most effective detection chemistry for the particular assay you are designing.

Overview

Detection of qPCR products may be achieved through the use of DNA binding fluorescent dyes. The dyes bind to any double-stranded DNA, which causes an increase in the intensity of their fluorescent emissions, as compared to their fluorescence in the unbound form.

As more PCR cycles are completed, copies of the original double-stranded DNA accumulate, allowing the dye to bind to additional double-stranded DNA. The fluorescence intensity continues to increase in proportion to the increase in the amount of PCR amplicons present.

DNA-binding dyes may be used for either singleplex or multiplex assays.

ADVANTAGES

Using a DNA-binding dye for your assay has several advantages. Overall assay design is much simpler, as there is no need to synthesize a probe for each target. This often means lower costs, increased convenience and decreased preparation time. They are easier to use, and appropriate for a wide range of applications.

DISADVANTAGES

The major disadvantage to using a dye for your assay is the non-specific binding of the dye to any double-stranded products. Since the dye will bind to any double-stranded DNA sequence, false positives are a concern, along with increased levels of background noise relative to your signal detection. In addition, some dyes cannot be used at high concentrations, as the dye will redistribute during the melt curve analysis.

Types of Dyes

SYBR Green

SYBR Green is the most commonly used fluorescent qPCR dye. It is a cyanine dye, and its high binding affinity for double-stranded DNA is due to the two positive charges it contains under typical PCR reaction conditions. After it is added to the sample, it binds by intercalating between the bases of the double-stranded DNA.

When the dye is bound to double-stranded DNA, blue light is absorbed by the complex and green light is emitted.

LIMITATIONS

Low fluorescence

PCR inhibition at high concentrations

Favored binding to GC-rich regions

Next Generation Dyes [Chai Green, EvaGreen, LC Green]

Although SYBR Green continues to be the most commonly used fluorescent dye for qPCR, there are several other fluorescent dye options available. These alternate dyes can present many of the same advantages as SYBR Green, but with fewer of the corresponding limitations associated with SYBR Green.

Optimal dye concentrations for these dyes are higher than for SYBR Green, maximizing the PCR signal. Saturation of the double-stranded DNA binding sites eliminates the potential for dye jumping during your melt curve acquisition, which makes these dyes highly suitable for applications involving high resolution melt curve (HRM) analysis, such as mutation or polymorphism detection.

WHY CHOOSE AN ALTERNATIVE TO SYBR GREEN?

Lower PCR inhibition

Provide better results for melt curve analysis

Can be used at saturating concentrations

Greatly increased fluorescent intensity

Lactic acid bacteria such as Lactobacillus and Pediococcus grow best in anaerobic (oxygen devoid) environments, so it is important to minimize the surface area as much as possible during the enrichment process. FastOrange B solution provides a limit of detection of 1 cell/sample in 48 hours when followed by qPCR. A volume of 50 mL is recommended for sensitive detection. Less volume of FastOrange B solution may be used, but this will result in decreased sensitivity.

Using the 50 mL FastOrange B Enrichment Bottles:

Add 50 mL of your sample to a 50 mL FastOrange B Enrichment Bottle using sterile technique and incubate it for 48 hours at room temperature (25 C +/- 2 C). Leave the bottle upright to minimize oxygen exposure and vent the sample to avoid pressure buildup.

Using the 240 mL FastOrange B Broth:

Aliquot your entire 240 mL FastOrange B Broth from the glass bottle into individual enrichment containers with your chosen volume. We recommend 50 mL for sensitive detection. Use sterile technique to transfer the solution.

Add your sample in a 1:1 ratio to one enrichment container, again using sterile technique. Incubate the enrichment container for 48 hours at room temperature (25 C +/- 2 C). Leave the bottle flat on its side and vent the sample to avoid pressure buildup.

After your sample has been enriched for 48 hours, proceed with analysis by qPCR.

All FastOrange solution and samples should be transferred utilizing sterile technique for enrichment. Prepare your workspace by clearing the area, sanitizing everything with 70% ethanol, and making sure you have all required materials in the immediate vicinity. Always use sterile pipettes and bottles.

1. Hold your container in the updraft of a strong open flame. If using plastic bottles, avoid direct contact with the flame to avoid melting. A Bunsen burner is ideal, but an alcohol lamp may also be used. Position the flame between your working hand and the containers.

2. Open the container with the solution to transfer at a 45 degree angle to minimize airborne contamination and hold the cap. If you must put the cap down, put it face down on a clean surface to minimize contamination from above. If using glass bottles, flame the openings by running them over the flame briefly. Do not directly flame plastic bottles or enrichment flasks.

3. Carefully transfer the desired quantity of solution in the vicinity of the open flame by pouring solution from the original bottle to the new container. Utilize a new container with graduation for correct measurement.

These bacteria require anaerobic (oxygen-devoid) growth enrichments, meaning there should be no air exposure for optimal growth. Anaerobic bacteria die easily in an aerobic environment, so it is very difficult to grow and detect these species even with enrichment and PCR. Your enrichment container should contain 50% enrichment solution and 50% sample and leave no room for air exposurei.e., if you utilize a 50 mL container, you should include 25 mL of FastOrange B solution and 25 mL of your sample.

To enrich, add 50% of sample and 50% of FastOrange B solution to your enrichment container to completely fill up the volume, topping off as necessary to ensure the solution reaches the rim of the container. Transfer your sample and FastOrange B solution using sterile technique. Incubate the container vertically at room temperature (25 C +/- 2 C) and do not vent to preserve the anaerobic environment. We recommend using sterile 50 mL Falcon or Eppendorf tubes or glass bottles, utilizing 25 mL of your sample and 25 mL FastOrange B Broth and topping off with sample to ensure that there is no air exposure in the container. The FastOrange B Enrichment Bottles are not recommended for this bacteria enrichment due to pressure buildup resulting from bacterial growth.

Enriching your samples before running direct PCR is recommended to improve sensitivity but requires extended incubation times. If enriching for L. acetotolerans, incubate the sample for at least 5 days. Sensitivity can increase by enriching up to 10 days. If enriching for Megasphaera and Pectinatus, incubate for a minimum of 7 days. Sensitivity can increase by enriching up to 12 days. Some anaerobic bacteria grow slowly and may have a long growth lag phase if exposed to oxygen during sampling or the enrichment process.

FastOrange Brett solution provides a limit of detection of 1 cell/sample in 72 hours when followed by qPCR. A volume of 40 mL is recommended for sensitive detection. Less volume of FastOrange Brett solution may be used, but this will result in decreased sensitivity. Brettanomyces grows best in anaerobic (oxygen devoid) environments, so it is important to minimize the surface area as much as possible during the enrichment process.

Using the 40 mL FastOrange Brett Enrichment Bottles:

Add 40 mL of your sample to the 40 mL FastOrange Brett Enrichment Bottle using sterile technique and incubate it for 72 hours at room temperature (25 C +/- 2 C). Leave the bottle upright to minimize oxygen exposure and vent the sample to avoid pressure buildup.

Using the 240 mL FastOrange Brett Broth:

Aliquot your entire 240 mL FastOrange B Broth from the glass bottle into individual enrichment containers with your chosen volume. We recommend 40 mL for sensitive detection. Use sterile technique to transfer the solution.

Add your sample in a 1:1 ratio to one enrichment container, again using sterile technique. Incubate the enrichment container for 48 hours at room temperature (25 C +/- 2 C). Leave the bottle flat on its side and vent the sample to avoid pressure buildup.

After your sample has been enriched for 72 hours, proceed with analysis by qPCR.

The Open qPCR software provides more detail for your kit runs by generating an Amplification Curve, which shows how many fluorescence units there are in the two fluorescence channels. Fluorescence indicates the presence of targeted DNA for each channel. To view the amplification curves for your experiment, select View Full Results at the bottom of your results screen. Make sure the Baseline Subtraction option at the top left of your curve is toggled on. You can toggle between the channels next to the Choose Channel option at the top right of your curve. Highlight wells below the curve to view individual well results.

The dual channel Open qPCR allows for multiple target detection. The target detected by Channel 1 is the beer spoiler DNA, which will be present in the Positive Control and any contaminated samples. Channel 2 detects the Internal Amplification Control in your negative control and clean beer samples. This channel functions as a quality control to detect potential PCR inhibition.

You can also analyze your results by looking at the Cqvalues in Channel 1 and Channel 2 in the table to the right of the Amplification Curve. The Cq value is inverse to the amount of target present in your reaction. A low Cq value indicates a high quantity of your target, while a high Cq value indicates a low target amount.

Channel 1: The positive control well should always amplify and generate a Cq Ch 1 value. The negative control should not amplify and should never generate a Cq Ch 1 value. Samples with trace amounts of beer spoiler will amplify and generate Cq Ch1 values; the more spoiler presence there is, the earlier the curve will amplify and the lower the Cq Ch1 value will be. Clean samples will have flat amplification lines and will not generate Cq Ch1 values. The curve below is from a successful run in which a customer ran a positive control, negative control, and two clean beer samples. The amplified blue curve is the positive control.

Channel 2: Wells that do not amplify in Channel 1 should amplify in Channel 2 and generate Cq Ch2 values. This includes the negative control and wells containing clean samples. If these wells do not generate Cq Ch2 values, your experiment experienced inhibition. Your positive control well may also sometimes generate a Cq Ch2 value. For cases when there is Channel 1 amplification, Channel 2 sometimes may not amplify; this is acceptable and does not indicate inhibition. In the curve below, the amplified curves are from the positive control, negative control, and beer spoiler wells, while the flat lines contain no tubes.

The enrichment process for acetic acid bacteria is similar to that of lactic acid bacteria, but optimal growth conditions vary. Unlike lactic acid bacteria, acetic acid bacteria grow best in aerobic (oxygen present) environments, so it is important to maximize surface area as much as possible during the enrichment process. FastOrange B solution provides a limit of detection of 1 cell/sample in 48 hours when followed by qPCR. A volume of 50 mL is recommended for sensitive detection. Less volume of FastOrange B solution may be used, but this will result in decreased sensitivity.

Using the 50 mL FastOrange B Enrichment Bottles:

Add 50 mL of your sample to a 50 mL FastOrange B Enrichment Bottle using sterile technique and incubate it for 48 hours at room temperature (25 C +/- 2 C). Leave the bottle flat on its side to maximize oxygen exposure and vent the sample to avoid pressure buildup.

Using the 240 mL FastOrange B Broth:

Aliquot your entire 240 mL FastOrange B Broth from the glass bottle into individual enrichment containers with your chosen volume. We recommend 50 mL for sensitive detection. Use sterile technique to transfer the solution.

Add your sample in a 1:1 ratio to one enrichment container, again using sterile technique. Incubate the enrichment container for 48 hours at room temperature (25 C +/- 2 C). Leave the bottle flat on its side and vent the sample to avoid pressure buildup.

After your sample has been enriched for 48 hours, proceed with analysis by qPCR.

Open qPCR does not require a ROX or any other reference dye.

Some instruments use a reference dye to normalize well-to-well differences. Because of Open qPCR's unique, solid-state optical architecture & calibration algorithms, no reference dye is required.

If you will be using a master mix containing a ROX reference dye, when calibrating your dual channel instrument, it is advisable to reconstitute your FAM + HEX calibrator set with your master mix, rather than Chai's reconstitution buffer solution.

The Open qPCR uses 0.1 mL low-profile PCR tubes. Both the tubes & caps should be optically clear. We recommend using Chai's PCR Tube & Cap Strips for the best results, which are also DNase, RNase, and PCR-inhibitor free.

What is PCR Efficiency?

PCR efficiency is defined as the percentage of target molecules that are copied per PCR cycle. Imagine a reaction tube that initially contains 100 target molecules. After one round of PCR, it now contains 190 target molecules. The PCR efficiency of this reaction is 90%, because 90% of the target molecules were amplified.

In any reaction, the ideal PCR efficiency is 100%. This indicates amplification of all of the target molecules with each successive cycle. However, a PCR efficiency of 100% is not always attainable. A PCR efficiency ranging from 90% to 110% is considered acceptable.

Why is PCR Efficiency Important?

If the underlying goal of the reaction is to quantify the total amount of target molecules in a given sample (quantitative PCR), a high PCR efficiency value is essential, as it is a reflection upon the accuracy of the data obtained from the reaction.

Calculating PCR Efficiency

PCR efficiency is calculated from the standard curve of an assay. A standard curve is created by preparing 5 individual serial dilutions of a control DNA template, each diluted by a factor of ten, against which the unknown target template can be measured. Each serial dilution is used to perform a separate real-time reaction. The subsequent Cq values are plotted and the standard curve is generated by fitting a linear line to the data points. The PCR efficiency can then be determined from the slope of the standard curve.

Low PCR Efficiency

The efficiency of a reaction may be negatively affected by a combination of a wide variety of factors. Some of the most common factors include:

Sample inhibition due to inhibitory compounds

Non-optimal PCR primer and/or probe design

Primers designed on a single nucleotide polymorphism (SNP) site

Incomplete DNAse treatment (if applicable)

Formation of primer dimers (PDs) or non-specific amplification

Dynamic range of the standard curve is not large enough

Incorrect sample dilutions

Pipetting errors across the standards and/or samples

Two major types of chemistries may be utilized to detect the products of your qPCR cycles, each with particular advantages. Understanding the differences between the two will ensure you choose the most effective detection chemistry for the particular assay you are designing.

Dye Chemistry Basics

Dyes that are used for qPCR molecule detection are small molecules with high binding affinity to any double-stranded DNA, including the original double-stranded DNA present in the sample and the new copies of double-stranded DNA being formed by the PCR cycles, using various binding methods. When the dye binds to double-stranded DNA, this causes an increase in the fluorescence, and the more DNA the dye binds to, the greater the intensity of the fluorescence. The intensity of the fluorescence by the dye increases proportionally to the increase in the amount of PCR products.

Dye Advantages & Disadvantages

The main advantage of using a dye for your assay is the reduced monetary and time investment required upfront, in order to get your assay up and running. Only a simple assay design is necessary, and no time needs to be spent synthesizing one or multiple probes. Additionally, double-stranded DNA binding dyes are useful if you plan to perform post-PCR analysis, such as high-resolution melting (HRM) analysis, However, since the dye will bind to any double-stranded DNA sequence, drawbacks that come with using a dye for your assay include possible false positive signals and overall decreased assay specificity.

+ Lower costs

+ Suitable for any double-stranded DNA sequence

+ No probe synthesis

+ HRM analysis

- Decreased specificity

Probe Chemistry Basics

The more specific method of detection involves the use of an oligonucleotide probe, which contains an additional fluorescent molecule, or fluorophore, attached to it. Two types of these fluorophores exist: reporter fluorophores and quencher fluorophores. The reporter fluorophore is found on the 5 end of the oligonucleotide and the quencher fluorophore is found on the 3 end of the oligonucleotide. When the reporter fluorophore initially absorbs energy from light, it enters an excited state. As it returns back down to its ground state, the energy is released as fluorescence, which is then transferred to the quencher fluorophore, a process known as Fluorescence Resonance Energy Transfer (FRET). This means fluorescence intensity emitted by the reporter fluorophore is greatly reduced when the quencher fluorophore is nearby, thereby making detection of the fluorescence difficult.

One of the most common types of qPCR probes are known as hydrolysis probes. When the reporter fluorophore of the hydrolysis probe becomes separated from the quencher fluorophore, such as via cleavage from DNA polymerase 5' to 3' exonuclease activity, the intensity of the fluorescence detected from the reporter dye will increase. As the amount of amplified product increases and additional probes are cleaved, the fluorescent signal measured will increase proportionally.

In addition to hydrolysis probes, there are other probe chemistries that produce the amplified fluorescent signal, such as Molecular Beacon probes and Scorpion primer-probes.

Probe Advantages & Disadvantages

In contrast to dye chemistry, one of the main benefits gained from utilizing probe chemistry is the increased specificity of your assay through the use of an oligonucleotide probe or primer. By using fluorophores attached to the specific oligonucleotide sequence, only the specific desired PCR products will be detected, and amplification of non-targeted products will be prevented.

The other major benefit is the possibility for optical multiplexing, or amplification and detection of multiple sequences within the same reaction. This capability is not achievable through the use of dye chemistry. Multiple probes are used, each labeled with a fluorophore. If the fluorescent signal generated by each fluorophore is detected in a different optical channel, detection of each distinct target sequence is possible. If fluorophores which all produce signals within the same optical channel are attached to multiple probes with different target sequences, multiple sequences may be detected at once, but differentiation between them is not possible.

+ Higher binding specificity

+ Multiple sequences may be amplified at once and distinguished by using different optical channels

- Different probes must be synthesized for different target sequences

Dye or Probe?

In general, choosing to use a dye for your assay may make more sense to you, if youre looking to meet these conditions:

More economical

Wide applicability

Design simplicity

Decreased set-up time

Post-PCR analysis

Conversely, using a probe for your assay may be the right choice, if the following criteria meet your needs:

Amplification of multiple target sequences

Detection of specific amplified products

Elimination of post-PCR processing

At the end of the day, thereisn'ta better choice when it comes to detection chemistry options. It depends on the priorities specific to your assay design, and the resources you have available. Factors such as cost, number of reactions and specificity requirements are important to consider when deciding which detection chemistry to use.

Probe qPCR detection chemistry involves the use of an oligonucleotide probe that contains attached fluorophores to allow for fluorescence detection of a specific amplified target sequence. Within this group exists three major classifications: primer-probes, hydrolysis probes and hybridization probes. The structures, along with the advantages and disadvantages of each, vary by type. Understanding these differences will allow you to choose the best probe for your qPCR assay.

Scorpions

Scorpion probes are classified as primer-probes, since they contain both a primer and a probe within the same molecule. Within this primer-probe classification, they are more specifically described as hairpin primer-probes.

When Should You Choose Scorpions?

Due to the unique unimolecular structure of Scorpions, these primer-probe molecules provide extremely fast detection of PCR products. Their structure also aids them in producing an excellent signal-to-noise ratio. These characteristics make Scorpions great options if your assay priorities are shorter reaction times, strong signals and enhanced discrimination. Additionally, Scorpions are a good choice when cost is an important consideration, as theydon'trequire purchasing both a primer and a probe.

Unimolecular Fast reaction kinetics

No primer-dimer formation

High specificity

Strong signals & minimal background noise

Multiplexing

Structure

This hairpin structure contains a reporter fluorophore at the 5 end of the single-stranded oligonucleotide and a quencher probe at the 3 end of the oligonucleotide. The loop portion of the hairpin contains a base sequence that is complementary to the sequence of the target DNA.

Additionally, a primer is attached to the hairpin structure. A hexathylene glycol (HEG) blocker attaches the 3 end of the hairpin to the 5 end of the primer. The HEG blocker ensures the primerwon'tbe copied by the polymerase.

Function

In this closed loop conformation, the intensity of fluorescence emitted by the reporter fluorophore is greatly reduced due to its quenching by the nearby quencher fluorophore, a process known as fluorescence resonance energy transfer (FRET). Following denaturation, the probe binds to the complementary sequence of the new DNA. When the probe binds to its complementary region of an amplicon, the hairpin structure opens and the reporter fluorophore separates from the quencher fluorophore. This separation of the reporter fluorophore from the quencher fluorophore allows the detection of the fluorescence signal from the reporter fluorophore to intensify. This increase in intensity occurs in proportion to the increase in the amount of amplicon.

Other primer-probes: Cyclicon, LUX

TaqMan

TaqMan probes are classified as hydrolysis probes. These probes are also classified as bimolecular, as the primers and probes are not attached within a single molecule.

When Should You Choose TaqMan?

TaqMan and other hydrolysis probes are normally your best option when cost is your top consideration, as they tend to be more cost-friendly. They are also tremendously dependable, making them a good option if youre looking to perform high-volume assays.

High specificity

Multiplexing

Easy probe design

Structure

TaqMan probes consist of a single-stranded oligonucleotide that is synthesized to bind to a specific single-stranded target DNA sequence. They contain a reporter fluorophore attached at the 5 end and a quencher fluorophore attached at the 3 end.

Function

The TaqMan probe anneals to the target region specified downstream of the primers used in the assay. As the DNA polymerase extends the primer and continues to synthesize the new strand, the reporter fluorophore of the probe becomes separated from the quencher fluorophore via cleavage of 5 to 3 Taq DNA polymerase exonuclease activity. The intensity of the fluorescence detected from the reporter dye is no longer quenched by the nearby presence of the quencher fluorophore via FRET. The probe is removed from the target strand as the polymerase continues to extend the strand. As the amount of amplified product increases and additional probes are cleaved, the fluorescent signal measured will increase proportionally.

Molecular Beacon

Molecular Beacon probes are a type of hybridization probe. They are single-stranded oligonucleotides containing a hairpin loop structure.

When Should You Choose Molecular Beacon?

If one of your top assay priorities is minimizing background noise and very high specificity, Molecular Beacon probes are an excellent option. In situations where your target sequence is highly uncommon, this can be very important.

Very high specificity

Multiplexing

Probe conservation

Structure

The structure of Molecular Beacon probes consists of four distinct regions. The loop portion contains the base pairs that are complementary to the target DNA sequence. The stem region of the hairpin contains two sequences, one at the 5 terminus of the probe and one at the 3 terminus of the probe, which are complementary to one another. Both ends of the stem region are attached to the loop region. The probes also contain a reporter fluorophore at the 5 end and a non-fluorescent quencher fluorophore at the 3 end.

Function

When the hairpin is in its closed conformation, the fluorescence emitted by the reporter fluorophore is quenched by the quencher fluorophore. Following denaturation of the target DNA and annealing of the primers, the hairpin probe unfolds and binds to the target sequence. The fluorescent emission from the reporter probe is no longer quenched, because the quencher fluorophore is no longer in such close proximity to the reporter probe. The fluorescent signal emitted by the reporter fluorophore is detected, and increases proportionally with the increase in PCR products produced. Since the hairpin loop will only unfold in response to an exact complementary sequence, these probes may be used when very high degrees of specificity are required.

Other hybridization probes: FRET, Eclipse