Magnetic beads from CleanNA have a Polystyrene core, covered by a magnetic layer and a caboxylate-modified layer. This layer is negatively charged and binds water. By adding EtOH, DNA will attach to the water. Due to the magnet layer, beads can be drawned to a magnet. This will allow you to remove the supernatant and wash your solution. When using the CleanNA kits automated, we recommend to use a ring magnet.
We recommend to use the lysis buffer from CleanNA. All buffers in the CleanNA kits are designed to optimize the outcome of the kits. When changing one of the buffers, for example the lysis buffer, the results can be lower than expected.
Yes, they can. Magnetic bead based isolations are very easy to automate, but can also be used manually.
The filter based method (also known as spin columns) has been the golden standard for Nucleic Acid isolation for a long time. Spin columns are easy to process, and prep time is relatively short. Yield and purity are consistent and high. So why use magnetic beads for your DNA or RNA isolation? Because spin columns are not easy to automate. There are a few manual steps involved when using spin columns that simply cannot be automated. When using magnetic beads, all steps can be automated using a liquid handler. You can either use a liquid handler already available on your lab, but you can also choose to go for a package-deal. The advantage of a package deal is that most suppliers will make sure that your protocols are pre-programmed on the liquid handler.
So what about the quality of my isolated DNA? Don’t worry about that! Compared to spin columns, DNA isolated with magnetic beads has the same quality.
An assay or application that is initially setup is often performed manually. This is very normal as there is a technician who is involved and little technical steps are performed in a very controlled environment (the technician’s eyes are on it). However, as the number of repeats increase or if the demand for the application increases a manual process is no longer the ideal solution. Automation is then the answer to make an application robust, reproducible and independent on technician skills. Automation is available in many forms (from a handheld repetition pipet up to a BioNex Hive platform). Once the step into automation is taken, the application can be considered to be of a higher throughput. The assay can vary and is not limited to rules. Any lab related process is able to be performed in high-throughput.
There is no straight forward definition of high-throughput. The definition is depending on many aspects such as the complexity of the assay, the number of samples and also the used labware and volume range. For compound screening in the traditional pharmaceutical companies, screening is done on 384 or 1536 well plates. Running 100 plates containing 1536 samples, results in 153.600 individual data points, safe to say that this is high-throughput. To compare, a bloodbank may run two completely automated DNA extractions on large volume blood (3 ml) and only be able to run 72 or 96 samples per day. This is also considered to be high-throughput.
As there is not a straight forward definition, high-throughput is more of a term used for an increased amount of actions to be performed.
A centrifuge needs to be in balance to operate properly. This is one of the first remarks made when an operator works with a centrifuge for the first time. There are numerous examples of accidents with centrifuges because of imbalanced loading. Luckily, modern technology allows for a tolerance in balance. This makes operating a centrifuge a lot easier for an operator.
Most automated centrifuges have two buckets to load plates. But not many automated systems are equipped with a weighing station. So, when working with plates with varying volumes in it, balancing out the centrifuge becomes a problematic step. Especially since a plate handler needs to have the same access point over and over again and moving a millimeter might already cause it to crash. The HiG centrifuges have a really high imbalance tolerance of up to 100 grams, without sacrificing into the speed (up to 5.000 g) eliminating the need for weighing a plate in most assays.
This depends on your application. A centrifugation step is expressed in a unit. This can be rounds per minute (rpm) or the relative centrifugal force (RCF) or times gravity (G). To convert from RPM into G, the following formula can be used:
g = (1.118*105) * R * S2
In this formula, R is radius of the centrifuge in centimeters and S is speed of the centrifuge in rpm.
Added to the centrifugation time is the time needed for acceleration to the needed maximum speed, and off course deceleration. In most assays, the centrifuge can actively decelerate, but in some assays the formed pallet is so sensitive that this is not advised. This may increase the centrifugation time tremendously. Normal acceleration and deceleration are done in less then 20 seconds. But when working with sensitive pallets, this may take up to 20 minutes since braking inside the centrifuge is completely turned off.
ROX is a fluorescent molecule that the real-time PCR system can detect when its present in the reaction. It’s used as a Passive reference dye for normalization of fluorescence signal across all of the PCR samples of the PCR thermo cycler including a baseline.
Rox is required when there is uneven illumination, sample variation and difference in quantity or condensation.
Its shadowing the reporter as a constant fluorescent and results in a precision of well data.
Adding ROX depends on which Real-time PCR instrument you’re using, you can find the exact recommended mix via the Real-time PCR selection table of Bioline
The SensiFAST and Sensimix kits has been developed for fast, highly-sensitive and reproducible qPCR and has been validated on commonly-used real-time PCR instruments and is designed for superior sensitivity and specificity with probe-detection technology, including TaqMan®, molecular beacon and Scorpions® probes. Consistently accurate detection of DNA and RNA targets from a broad range of sample types and Excellent efficiency for improved multiplexing
• Sensifast includes The use of an antibody-mediated hot-start DNA polymerase which minimizes amplification from primer-dimers, thereby improving assay specificity and sensitivity. Sensimix includes a Covalently-modified hot-start promotes highly-specific amplification
• Sensifast is the latest version and improved Reproducible. Faster accurate results in as little as 30 minutes.
1 step kit involves including the reverse transcriptase step in the same tube as the PCR reaction and the 2-step kit involves creating cDNA first.
Using gene specific primers, one-step real-time RT-PCR such as the One-Step kits offer a quick and simple method to detect mRNA and so are useful when analyzing a few genes over a large number of samples as less pipetting and sample manipulation reduces variation and potential contamination. However reaction conditions needed to support both the RT and PCR may not be optimal for either reaction and it is not possible to archive the cDNA produced during the reverse transcription reaction.
This method is quick to set up and makes processing multiple RNA samples easy (especially when using liquid handling robotics), when you are amplifying only a few genes of interest. It is therefore ideal for high throughput screening laboratories where only a few assays are run repeatedly, using well-established reaction conditions, with the added advantage that multiplex PCR of the gene of interest and control genes can be done in single well.
Two-step real-time RT-PCR, offers a truly accurate determination of mRNA and is useful when analyzing a large number of transcripts over a few samples. SensiFAST™ kits have flexibility in the priming strategy, allowing for oligo-dT, random primers or gene specific primers and are generally more sensitive than one-step as the RT and PCR occur separately and can be optimized individually. Also, the cDNA produced is more stable than the initial RNA sample and can be more easily archived for future use.
With two-step real-time PCR, the use of several tubes means that it is more time consuming and less adaptable to liquid handling robotics and so more difficult to adopt for high throughput screening assays. The use of several tubes and pipetting steps also exposes the reaction to a greater risk of DNA contamination
– Accurate representation of target copy number
– Simple and rapid Fewer pipetting steps (reducing possible errors and contamination)
– Best option for high-throughput screening
– Best method when only a few assays are run repeatedly
– Multiplex PCR of gene of interest and control can be done in single well, from same RNA sample
– Usually less sensitive as it is impossible to optimize the two reactions separately
– Difficult to troubleshoot RT step
– No stock of cDNA
Before choosing you should consider the best one for your application, these include the ease of use and cost of reaction to the resulting yield and sequence representation.
In most PCR and qPCR reactions Thermus aquaticus Polymerase (TAQ polymerase) is used as a polymerase.
The main reason to us a hot start Polymerase (also known as a ‘Hot Start PCR’) in your PCR reaction is to avoid unspecific amplification. Most DNA polymerases work best at a temperature between 68 and 72°C. In some cases, an enzyme can become slightly active below these temperatures and this will cause unspecific binding, leading to unspecific amplification. A hot start PCR will reduce the nonspecific amplification significantly.
Besides avoiding the unspecific amplification, a hot start Polymerase will reduce the formation of primer dimers. Primer dimers can also be prevented by lowering the concentration of Primers, Magnesium Chloride or Nucleotides.
Last but not least, when using a hot start polymerase often product yields will increase!
So why not always use a hot start polymerase? Well, it has a disadvantage as well.. The re-activation time during the denaturation stage is increased for activation of the enzyme. This increased heating time could damage your DNA. Studies have also shown that using a hot start PCR can cause issues when amplifying long strands of DNA.
PCR measures the end-point, for example using an Agarose Gel. Nowadays people often use a quantification system like the DS-11, since this will give detailed information about the concentration and purity of your PCR product.
When performing a real-time (or qPCR) reaction, DNA can be measured during each cycle, allowing you to follow the amount of DNA present in your reaction tube.
PCR (polymerase chain reaction) is a technique to make copies of a DNA segment. It requires a thermocycler, a DNA polymerase, Nucleotides, Primers and off course your DNA template. After adding all ingredients together for your PCR reaction, the thermocycler will do the rest. During a number of cycles, it will heat up and cool down. There are three main stages during each PCR cycle:
Denaturing: The double stranded DNA is heated to separate it into two single strands
Annealing: The temperature is lowered to enable the DNA primers to attach to the DNA template
Extending: The temperature is raised and the new strand of DNA is made by the DNA polymerase, using the Nucletides to build the strand.
After each PCR cycle, the number of DNA strands are doubled.
The exact temperatures of each stage depend on the Primers, DNA Fragment and DNA Polymerase being used. In general the temperature of the denaturing stage is 94-95°C, annealing stage is between 50-65°C and extending stage is around 72°C.
260/280 ratio is a potential measurement for the detection and quantify of DNA / RNA.
The ratio of absorbance at 260 nm and 280 nm is used to assess the purity of DNA and RNA extractions. That is because Nucleic acids shows the highest absorbance generation/highest UV radiation absorbance at 260 nm and maxima at 280 nm.
It can also be used to check whether its contaminated.
A 260/280 ratio between 1.8 – 2.0 is generally accepted as “pure” DNA/RNA.
If the ratio is below 1.8 then it can indicate the presentence of proteins or phenyl contamination.
More and more people are using home made beads, but why would you choose the one or the other. Looking at the results of the home made beads vs the commercial available beads there is not a big difference, both beads contribute to clean DNA. A big difference between the home made beads and commercial beads is that the commercial beads have quality controls during the process, which contribute to reproducibility of the beads, when looking at home made beads these controls would have to be down by yourself. Home made beads are cheaper, but more labor intensive and possibility of a larger standard deviation in the reproducibility of the beads.
Is it possible to make home made beads for RNA? This is possible as long as you work in a RNase free environment and with chemicals that are produced and kept RNase free. So it is possible but it is also possible to order commercial beads that are produced RNase free.
For next generation sequencing (NGS) people are using double size selection. But why is this useful?
Size selection has one of the greatest impact on quality of results, when working with size that is not interesting, can waste sequencing capacity on low molecular weight material thinking of adapter-dimers or primer-dimers. Size selection can boost sequencing efficiency and save money. When using size selection to remove smaller or large fragments, which allows the sequencers to focus on the DNA fragments most interesting.
Let’s begin with what is a clean-up, a clean-up is another word for purifying your DNA. This can be after an isolation of DNA where the solution could still contain inhibitors or can be used after a PCR reaction, to remove the remaining primers, Taq, dNTP’s and MG ion, these can be seen as impurities for downstream application. After DNA isolation the inhibitors can stop or slow down your PCR products or qPCR which can give a fault to your results. If you are planning to run a sequence thinking off Sanger or Next Generation Sequencing purified DNA is necessary to not interfere with your results.
The clean-up after PCR can be done with ethanol precipitation, enzymatic cleanup, column or bead based purification. Ethanol precipitation is cost effective, but also labor intensive. With enzymatic clean up you are not limited by starting material but has to sit with a certain temperature. Column based is also an effective method but there is a possibility that you loose a part of your yield due to that the DNA sticks to the filter. Another method is with magnetic beads, for this you are limited by the starting materials but easy to use. If you are interested in using magnetic beads look at https://gcbiotech.com/product/cleanna-ngs/
During sanger sequencing many copies of target DNA region are made. For this the next ingredients are need: your template DNA, DNA polymerase enzyme, a primer, the four DNA nucleotides (dATP, dTTP, dCTP, dGTP) and dideoxy (ddATP, ddTTP, ddCTP, ddGTP) also know as the dye.
By adding a high concentration of dye is added, this can give blobs on the sanger sequencing results, for this you can choose to add magnetic beads to remove the extra dye.
Sequencing is a process to determine the sequence nucleotides (A, T, C and G’s) in a piece of DNA.
NGS is the faster form of sanger sequencing which can do massively samples in parallel while sanger sequencing provides one forward and reverse read. Sanger sequencing has a 99% accuracy and is the golden standard but takes more time to sequencing multiple samples. Sanger sequencing is often used as a confirmation of the NGS run.
The use of the type of tip (carbon or transparent) is depending on the type of pipetting technology (see “what is the difference between system liquid and air displacement”). The technology determines the technique used for liquid level detection in the system.
Black carbon tips are used when the liquid level detection is based upon conductivity. The ions in the liquid will cause a signal through the tip when the tip touches the liquid. This signal tells the system the liquid level has been found. Systems operating on system liquid will always use the black carbon tips.
Liquid level detection can also be done using transparent tips. The basic principle is different. There will be a pressure or flow sensor in the channel. When the tip touches the liquid, the pressure in the channel changes slightly.
The advantage of the use of transparent tips is often in the price. If the application does require the use of a lot of tips, or if tips cannot be washed, this advantage increases tremendously over the lifetime of the system.
The difference between air displacement and system liquid is in the way the pipetting is performed. Basically when having system liquid, the aspiration and dispense motion is created by moving a column of fluid (often water) through the tubing with a pump. An airgap between the system liquid and the aspirated fluid makes sure that the system fluid will not be contaminated. Off course the system fluid does not travel through the tip.
Air displacement does exactly the same, but instead of moving system fluid, it moves air. The upside of air displacement is that you will need a lot less tubing in the system, it does not require any additional fluids to be filled and maintained and so on. Especially in Life Sciences the use of air displacement over system fluid gives advantages.
Some low volume dispensing techniques, such as the BioNex Nanodrop, use pressurized system liquid. The advantage of using pressurized system liquid is an increased accuracy in the dispense enabling the system to dispense volumes of 100 nl very accurate. The airgap ass mentioned above will then also play a role in the accuracy of the dispense.
Contamination occurs when droplets are formed at the end of a tip, when aerosols are formed during the application and so on.
The droplets are a problem that can and must be prevented. The way to do this is by aspirating a transport airgap when traveling across the deck with liquid in the tip. This is a standard point in optimization as the transport airgap varies in size depending on a few factors (type of liquid in the tip, volume in the tip, volume to be dispensed etc.).
Aerosol formation is harder to prevent, especially in very sensitive applications. However, most liquid handlers can be equipped with a HEPA filtration unit, UV lights and other additional features to keep the system as clean as possible.
In a system, contamination is formed when the sample is aspirated into the system. A liquid handler using disposable tips will only aspirate samples in that tip. This ensures no samples in the system itself. From a manual experience, many users are persuaded to use filtered tips, also on a liquid handler. However, opposed to working manually, if the system is optimized and used correctly, there is no need for filtered tips for 99% of the applications.
In systems without a disposable tip, needles or nozzles are used to aspirate and dispense. There is a higher risk of contamination using this technique, but it does make washing easier. In regards to using nozzles, the main source of contamination is found in all connection points and especially in valves if not washed correctly.
Programming a liquid handler is a specialized task. Each liquid handler has its own software. In the beginning of automation, these software packages were very basic and required a lot of scripting knowledge. Nowadays, most liquid handlers have a ‘drag-and-drop’ software interface, decreasing the complexity of programming tremendously. Adding a basic application is often a reasonable task for a champion user.
If the application requires scheduling, parallel processing, or complex calculations, it may be wise to consult your supplier before you start programming.
GC biotech tries to enable everybody to at least use the liquid handler for most daily tasks. We do this by decreasing the complexity of the daily use as much as possible in our programming approach and with additional software skills. During the installation of each system, a user training and programming training can be provided.
Arranging the deck layout is not bound to any set rule. The application itself and the demands on the application will determine the layout for a big part.
Automation of any laboratory workflow is based upon many demands. One of the most often heard demands is to increase the speed, but this is a wrong assumption. A technician is very often faster than a liquid handler. The advantage of automation in this case is that the hands-on time of the application is decreased drastically, enabling the option to perform other tasks such as data interpretation.
Other reasons for automating any workflow, are increasing throughput, decreasing errors and increasing reproducibility. These are all valid reasons to start looking into automation.
A liquid handler has a higher accuracy, precision and especially reproducibility over a technician. It also never needs a break (except when it’s due for maintenance) and it is not hungover after the annual Christmas party. As good as this sounds, liquid handlers have many limitations that may cause a user to not choose for automation. These are all the little tricks and adaptations that a manual pipet makes, like swirling around a crystalized particle.
When automation will become a serious option, it will not just take the place of the technician. It very often has an impact on the entire workflow, especially if there is no prior experience with automation of similar processes. A lot of the preparation work is invested in looking at the workflow and the goals of the process. Together with an applications specialist the workflow, the goals and the criteria needed for successfully automating the process are retrieved. This will help us understand your needs, and help you understand what we will do for you.