Product FAQ

If I do not use all the wells in the filter plate of the PureLink Pro 96 Purification System, can I use those wells at another time?

Answer

Yes. Keep the unused wells covered with Foil Tape. Please note that extra Foil Tape and Receiver Plates will be needed.

Answer Id: E3805

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Product FAQ

What filter plate do you recommend be used with the Luminex reagents?

Answer

We recommend the Millipore filter plate (Cat. No. MSBVN1250). This plate has a 1.2 micron pore size. Our buffer kits and ready-made multiplex assays also include one of these plates.

Answer Id: E5151

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Product FAQ

My filter plate is leaking. What should I do?

Answer

Here are some possible causes and solutions for this problem:

Placing the plate on an absorbent material both at the bench and on the shaker (which results in wicking when there is contact between the absorbent material and the opening on the bottom of the well funnel). Rest the plate currently in use on top of a second filter plate so that the filter plate currently in use only comes into contact with a nonabsorbent surface. This will also help locate any potential leaks.
Prior to incubations, use a Kimwipe tissue and lightly press up on each well to dry off the bottom of the plate. Putting any vertical pressure on the plate during the incubations (using either tape or clamps).
Use clamps that fit around the sides of the filter plate to secure the plate to the shaker during incubations. Subjecting the plate to vacuum pressure greater than 5 mm Hg, even for a moment, which either tears the filter membrane or creates an opening that exceeds the design limits for volume retention.
Prevent any vacuum surge by opening and adjusting the vacuum on the manifold before placing the plate on the manifold surface and checking the vacuum with a test plate (i.e., not the plate used for the assay). For our assays we recommend the setting not exceed 5 mm Hg. Puncturing the filter membrane with pipet tips, by inserting the tips all the way into the wells during reagent loading.
Pipet solutions along the sides of the wells, rather than deep into the wells. Separating the soft (opaque) plastic bottom from the hard (clear) plastic top of the filter plate before starting, which compromises the integrity of the seal even if the pieces appear to fit back together properly. If you accidentally separate these two layers of the plate, the plate should be discarded and a fresh plate used instead.

Answer Id: E12654

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Product FAQ

My filter plate has clogged wells. Do you have any suggestions?

Answer

If wells are clogged after the first incubation with the standards:

When washing a partial plate, the vacuum may not be complete because air is being drawn through the empty wells. We recommend covering the plate with Parafilm film to create a seal, which will in turn raise the pressure high enough to empty the wells. If there is a small clog, it may be difficult to see it even though it is enough to prevent aspiration. For this, we suggest using the tip of a 15 mL conical tube to gently rub against the tip of the well bottom opening, going from left to right. This will dislodge small clogs only. Other ways to unclog a well are to apply pressure from above the well using a gloved finger or thumb with an absorbent paper towel under the plate, or unplugging the drain hole using a large syringe needle.

Answer Id: E12656

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Manual / Product Insert

96 well Filter Plate

Version: JAN.17
Catalog #

Product FAQ

How do I prevent the Luminex filter plates from leaking?

Answer

Here are the most common causes of Luminex filter plate leakage:

(1) The wells are porous and designed to permit flow. If the plate is placed on a wet surface, liquid can potentially enter the wells, altering the volume of liquid in the wells, along with the concentrations of assay constituents. If the plate is placed on an absorbent surface, liquid will be lost from the wells due to wicking action.
Solution: Make sure that the plates are placed only on clean, dry, non-absorbent surfaces during loading and incubations. After washing, make sure to blot residual liquid droplets from the bottom of the plate with clean paper towels.

(2) Putting any vertical pressure on the plate during the incubations (using either tape or clamps).
Solution: Use clamps that fit around the sides of the filter plate to secure the plate to the shaker during incubations.

(3) Subjecting the plate to excessive vacuum, even for a moment, can potentially tear the membrane.
Solution: When evaluating a new vacuum manifold for use with these reagents, the vacuum setting should be adjusted until about 3 seconds are required to empty the wells of 0.2 mL solution. Vacuum surge should be prevented by turning on the vacuum manifold before placing the plate on the surface.

(4) The membrane is punctured by pipette tips as liquids are added.
Solution: Pipet solutions along the sides of the wells, rather than directly into the wells, to prevent puncture of the membrane.

(5) Separating the clear top of the filter plate from the opaque bottom portion compromises the seal.
Solution: If you accidentally separate these two layers of the plate, the plate should be discarded and a fresh plate used instead.

Answer Id: E5150

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Product FAQ

Are the PureLink 96/384 systems compatible with the Qiagen robotic system?

Answer

The original QiaRobot does not allow modifications to its protocol; however, more recent models are more flexible. Contact Technical Support to learn about robots that are compatible with the system.

Answer Id: E3803

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Product FAQ

What size plasmids can be isolated using the PureLink 96/384 system?

Answer

Plasmids up to 28 kb have been isolated and sequenced.

Answer Id: E3806

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Product FAQ

Can I use DNA purified using the PureLink 96/384 Plasmid Purification System in transfection protocols?

Answer

No. While the DNA is ideal for automated fluorescent sequencing, PCR, and restriction enzyme digestion, it is not compatible for eukaryotic transfection.

Answer Id: E3807

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Product FAQ

Can I use a low-copy number plasmid for the PureLink 96/384 plasmid purification system?

Answer

No. This product is not recommended for use with low-copy number plasmids when the plasmid DNA is for use in automated fluorescent DNA sequencing. Low-copy number plasmids can be used with this system when the samples are for use in PCR amplification. However, keep in mind that the yields from a low-copy plasmid may not be readily visible on a gel.

Answer Id: E3808

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Product FAQ

Are there any general recommendations or points to consider when setting up a Luminex assay?

Answer

TIP #1: Use a bunch of paper towels to blot the plates after each wash step. After removing the plate from the vacuum manifold, there are usually droplets of liquid still present on the bottom of the plate. By blotting you remove this, eliminating the possibility that the droplet will act as a wick when in contact with another surface and leak volume out of wells.

TIP #2: Shake the plates as fast as possible to allow for maximum vortex without spillage. The beads are conjugated with monoclonal antibodies over their entire surface area. Think of each bead as being the same as one well in an ELISA plate. Since the beads are heavy and sticky, they sink to the bottom of the well if left in standing solution and then clump together. Once they are clumped together on the plate, you cannot separate them and you will have aggregation problems during the plate read. To avoid this, after sonicating them, we shake the beads on the plate at a speed that allows for maximum vortex without spillage.

TIP #3: As a rule, the strength of the vacuum filtration setup should be such that it takes three seconds to empty each well of wash solution. Since there is extraordinary variation in the strength of vacuum lines and pumps, it is safer for you to use this guideline rather than a value. Since the filter membrane is relatively fragile, applying too much pressure, even for an instant can cause tears and subsequent bead loss (vacuum flow should be established before placing the plate on the manifold) and DO NOT EXCEED 5 mm of Hg . The three-second-count rule is an easy and safe guideline.

TIP #4: Spin all samples down prior to loading. Another feature about filter plates is the possibility for them to become clogged. This happens when the sample has visible precipitate or aggregated matter, which most commonly occurs for serum and plasma samples (based on collection techniques). While we typically do not recommend spinning down TCS samples, we have seen this sample type clog a plate as well. Since clogs cannot be resolved once they occur, meaning a loss of data for the well, it is better to prevent the clog by spinning the samples down prior to loading. Generally, a 1 min high-speed spin is sufficient for most samples.

TIP #5: Preload samples onto a nonbinding/low binding microtiter plate to allow for easy transfer and quick loading in the assay. A 96-well plate represents a large number of samples, particularly when you are not running duplicates. Because binding begins immediately once a sample is added to a well, the difference in incubation time between the first and last well can be substantial. This can have an effect on the sample values, along with exposing the beads to excessive amounts of light during loading. The easiest way to avoid this, is to preload an aliquot of samples onto a microtiter plate and then simply transfer them with a multi-channel pipettor at the appropriate point in the assay.

TIP #6: If a well is clogged, use the bottom of a 15 mL conical tube to very gently press the bottom tip of the well from left to right once before attempting to apply vacuum again. A clogged well only occurs following the sample incubation step. When this happens, it is important that you do not raise the vacuum pressure, since this will risk tearing the other wells. Instead, use the tip of a 15 mL conical tube to gently press the bottom tip of the well funnel from left to right once. Try to aspirate the sample again, and if that does not work repeat the gentle pressing. After two or three attempts, you can consider the well permanently clogged and the data lost. It is not possible to dissolve major clogs like this with high detergent, because the detergent will also disrupt the antibody binding. It is also not possible to manually empty and wash the well from the top end, since beads will be removed each time until the point where there are none left to be analyzed.

Answer Id: E5139

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Product FAQ

What is the best method for quantitating the DNA purified from the PureLink 96 Plasmid Purification System?

Answer

For high-throughput quantitation, a method that uses PicoGreen or Hoechst Dye 33258 is recommended. Contact Thermo Fisher Scientific Technical Support for a protocol. The Quant-iT DNA Assay Kit, Broad Range is a fluorescence-based assay that can be used for accurate quantitation of DNA as well. A Lysis Buffer component makes A260/A280 ratios unreliable. However, the component does not affect DNA sequencing, PCR, or restriction enzyme digestion.

Answer Id: E3802

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Product FAQ

How do I process urine sample for use on the Luminex assay platform?

Answer

Use only freshly collected urine samples. Dilute 2-fold with the Assay Diluent provided in the kit. The final dilution of the sample will be 4-fold, and all results should be multiplied by 4. As needed, clarify samples by centrifugation (14,000 rpm for 10 min) and/or filter them prior to analysis to prevent clogging of the filter plates and/or probe.

Answer Id: E12644

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Product FAQ

How do I process synovial fluid on the Luminex assay platform?

Answer

Collect synovial fluid into non-heparinized tubes and spin at 1,000 x g for 10 min within 30 min of sample collection. The acellular portion of synovial fluid should be stored at -80 degrees C before subsequent analysis. All samples need to be clarified by centrifugation (14,000 rpm for 10 min) and/or filtered prior to analysis to prevent clogging of the filter plates. Dilute samples 1:1 with Assay Diluent prior to addition to the assay. Reference: Raza K et al. (2005) Arthritis Research &Therapy 7(4): R784-R795.

Answer Id: E12645

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Product FAQ

How do I process cell lysates (extract cellular proteins) for the Luminex assay platform?

Answer

The protocols mentioned below have been applied to several human and mouse cell lines. You should optimize the cell extraction procedures for your own applications.

Cell Lysis Procedure

Non-adherent cells:
Pellet cells by low-speed centrifugation. Remove the medium from the pellet, and wash twice with ice-cold PBS. Remove the PBS, and resuspend the cell pellet in cell lysis buffer (recommended cell lysate concentration is 2-5 mg/mL). Incubate 15 min on ice with occasional vortexing. Transfer the lysate to a microcentrifuge tube and centrifuge at 14,000 rpm for 10 min at 2-8 degrees C. Aliquot the cleared lysate into clean microcentrifuge tubes, and determine total protein concentration.

Adherent cells:
Remove the tissue culture medium from the cells, and wash twice with ice-cold PBS. Remove the PBS, add cell lysis buffer (recommended cell lysate concentration is 2-5 µg/mL), and incubate 15 min on ice. Collect the cell lysate and transfer it to a microcentrifuge tube. Centrifuge at 14,000 rpm for 10 min at 2-8 degrees C. Aliquot the cleared lysate into clean microcentrifuge tubes, and determine total protein concentration. Lysates should be frozen and stored at -80 degrees C or analyzed shortly after collection. Avoid multiple freeze-thaw cycles of frozen samples. Thaw completely, mix well, and clarify by centrifugation (14,000 rpm for 5 min) prior to analysis to prevent clogging of the filter plates.

Recommended Cell Lysis Buffer:
NP40 Cell Lysis Buffer (Cat. No. FNN0021) Note: Lysates prepared with NP40 Lysis Buffer must be diluted at least 5-fold prior to running the assay. The recipe to make the buffer is as follows: 50 mM Tris, pH 7.4, 50 mM NaF, 260 mM NaCl, 1 mM Na3VO4, 5 mM EDTA, 0.02% NaN3, 1% Nonidet P40. The NP40 Cell Lysis Buffer (without protease inhibitor cocktail and PMSF) is stable for 2-3 weeks at 2-8 degrees C, or for 6 months when stored in aliquots at -20 degrees C.

Add fresh to the NP40 Cell Lysis Buffer just before use:
1 mM PMSF (stock 0.3 M in DMSO)
Protease inhibitor cocktail (Sigma, Cat. No. P-2714)
Alternate Cell Extraction Buffer
Cell Extraction Buffer (Cat. No. FNN0011) Note: Lysates prepared with Cell Extraction Buffer must be diluted at least 10-fold prior to running the assay. Or 10 mM Tris, pH 7.4 2 mM Na3VO4 100 mM NaCl 1% Triton X-100 1 mM EDTA 10% glycerol 1 mM EGTA 0.1% SDS 1 mM NaF 0.5% deoxycholate 20 mM Na4P2O7 The Cell Extraction Buffer (without protease inhibitor cocktail and PMSF) is stable for 2-3 weeks at 2-8 degrees C, or for 6 months when stored in aliquots at -20 degrees C. Add fresh to the Cell Extraction Buffer just before use: 1 mM PMSF (stock 0.3 M in DMSO) Protease inhibitor cocktail (Sigma, Cat. No. P-2714)

Answer Id: E12651

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