Pipets cleaning

Three important precautions are needed when working with pipets and volumetric flasks. First, the volume delivered by a pipet or contained by a volumetric flask assumes that the glassware is clean. Dirt and grease on the inner glass surface prevents liquids from draining evenly, leaving droplets of the liquid on the container s walls. For a pipet this means that the delivered volume is less than the calibrated volume, whereas drops of liquid above the calibration mark mean that a volumetric flask contains more than its calibrated volume. Commercially available cleaning solutions can be used to clean pipets and volumetric flasks. 

Make sure that there are no air bubbles. I usually place 40-50 pL PCR reaction mix on the top of the coverslips. Invert one slide at a time, touch it to the drop of reaction mix, and quickly lift the slide. The coverslip should remain adhered to the slide because of surface tension. Flip the slide, center the coverslip with a clean pipet tip, and gently remove the air bubbles. 

Proper reagent care can reduce problems stemming from contamination, heat or excessive light exposure. Reagent contamination can be avoided by the use of clean pipet tips. Prompt return of reagents to proper storage conditions will prolong their shelf life. 

Draw the cabbage juice indicator solution into a clean pipet, and add 5 drops to each of the solutions in wells H1-H8. Stir the solution in each well with a clean toothpick. 

Use a clean pipet to carefully add a drop of the standard NaOH solution to the solution in well Al, and stir with a toothpick. Pause for about 30 seconds and look down through the well for evidence of a persistent, light pink, phenolphthalein color that indicates the endpoint of the titration. Repeat this process with each drop until the endpoint is reached. Record the num-... 

Lay out the test strip on the bench. Using a clean pipet tip each time, remove a l-pL aliquot from each collected fraction and spot it into the center of the correspondingly numbered square on the test strip. 

All necessary materials should then be placed in the cabinets, segregating clean items, cultures, and receptacles for contaminated items. The worker should organize the materials so that contaminated materials do not need to pass over clean items (remember, the air flow direction is downward). A good layout of materials would place cultures, clean pipets, flasks, etc. toward the front on either side of the work area, and a discard tray for culture plates and pipets towards the rear of the work space, as shown in Figure 9.9. Avoid placing items within ten centimeters of the cabinet front. The cabinet must not be overloaded with materials. Overloading will disrupt the air stream and interfere with proper cabinet performance. 

Micro-pipetting instruments such as the "Eppendorf or "Oxford pipettors with disposable plastic cone tips are customarily employed to dispense the liquid samples into electrothermal atomizers. Sampling problems which are associated with the use of these pipettors are among the troublesome aspects of electrothermal atomic absorption spectrometry (67,75). The plastic cone-tips are frequently contaminated with metals, and they should invariably be cleaned before use by soaking in dilute "ultra pure nitric acid, followed by multiple rinses with demineralized water which has been distilled in a quartz still. 

Even inside the controlled conditions of a research laboratory, analyzing clean and standardized test samples PCR procedures requires careful quality control, taking into consideration differences in sample preparation, variation in pipetting, differences in reaction tube thickness, poor calibration or instability of the thermal cycler, and reagent quality. 

The advantages of this screen filter, as cited by the authors, consist of its reusability, its standard 96-well format size, and its ability to be used whenever sample transfer or pipetting is needed. After usage, the screen filter can be easily cleaned by rinsing with water and methanol and additionally, by ultrasonication in water or methanol or other appropriate solution. The filter can be inserted into a plasma storage plate before sample transfer by the Tomtec Quadra used by the authors for automation. 

Again, with a disposable pipet, carefully add clean eluent until you re sure all the sample is stuck on the adsorbant, and none of the sand. 

The use of clean glassware is of utmost importance when doing a chemical analysis. In addition to the obvious need of keeping the solution free of contaminants, the walls of the vessels, particularly the transfer vessels (burets and pipets), must be cleaned so that the solution will flow freely and not bead up on the wall as the transfer is performed. If the solution beads up, it is obvious that the pipet or buret is not delivering the volume of solution intended. It also means that there is a greasy him on the wall that could introduce contaminants. The analyst should examine, clean, and reexamine his or her glassware in advance so that the free how of solution down the inside of the glassware can be observed. For the volumetric flask, at least the neck must be cleaned in this manner so as to ensure a well-formed meniscus. 

Pipets pose a special problem. Brushes cannot be used because of the shape of some pipets and the narrowness of the openings. If soap is to be used, one must resort to soaking with a warm soapy water solution for a period of time proportional to the severity of the particular cleaning problem. Commercial soaking and washing units are available for this latter technique. Soap tablets are manufactured for such units and are easy to use. 

Give both your acid and base solutions one final shake at this point to ensure their homogeneity. Rinse the buret with 5 to 10 mL of NaOH twice, and then fill it to the top. Open the stopcock wide open to force trapped air bubbles from the stopcock and tip. Allow this excess solution to drain into a waste flask. Bring the bottom of the meniscus to the 0.00-mL line. Using a clean 25-mL pipet (volumetric), carefully place 25.00 mL of the acid solution into each of the three flasks. Add three drops of phenolphthalein indicator to each of the three flasks. 

Prepare five identical dilutions of the stock solution prepared in step 1 by diluting 10 mL of the solution to 100 mL. Use a clean 10-mL volumetric pipet and a clean 100-mL volumetric flask, and make the measurement and transfer as carefully as possible (see Table 4.1). Label each as 2/1, 2/2, etc., to indicate that they are solutions 1 to 5 prepared in step 2. 

Pipet 100.0-mL aliquots of the water sample (an aliquot is a portion of a solution) into each of three clean 500-mL Erlenmeyer flasks. Add 5.0 mL (graduated cylinder) of the pH =10 buffer and three drops of EBT indicator to each flask. 

Prepare 100 mL of a 100 ppm Fe stock solution from the available 1000 ppm stock solution. Use a clean 100-mL volumetric flask and a clean volumetric pipet. Shake well. 

For each the calibration standard, unknown, control sample, and distilled water for the blank, pipet 50.00 mL into a clean, dry 125 mL Erlenmeyer flask. Add 8.0 mL of the combined reagent and mix thoroughly. After at least 10 min, but no more than 30 min, measure the absorbance of each at 880 nm. 

Prepare samples (maximum of two) by grinding and weighing 5 g of each into 500-mL Erlenmeyer flasks. Add 50 mL of the HC1 solution prepared in step 1 to each. Bring each to a boil on a hot plate, and then simmer for 5 min. Cool and transfer the supernatents for each to separate 100-mL volumetric flasks. Dilute to the mark with distilled water and shake. Filter each through Whatman 1 filter paper into dry 250-mL Erlenmeyer flasks. Pipet 1 mL of each of these extracts into other clean 100-mL volumetric flasks, and dilute to the mark with water. Save the original extracts in case more dilutions are needed. 

Exogenous sources such as a person s hair or skin, doorknobs, laboratory benches, dust, reagents, thermal cyclers, and pipet tips are some of the common sources of DNA contamination. Ideally, a laminar air flow bench with filtered air provides a clean, dust-free environment. Sample preparation should be done in a separate room or area. The addition of sample to the PCR reaction mixture in the... 

A known volume of KBr solution is pipetted into a clean beaker and a freshly polished silver electrode is then immersed in this solution. After a potentiometric titration with AgNOj solution at 298 K, the electrode potential at the equivalence point (EAg+,Ag) is determined to be 0.441 V. What is for the pale yellow AgBr that is formed ... 

Pipet 16 mL of the clarified serum or ascites into each of six clean 50-mL polycarbonate tubes. Add 16 mL of cold, saturated ammonium sulfate solution and stir gently with a pipet (see Note 6). 

Pipet 100 pL of the cell suspension onto treated slides, spread using the edge of a clean slide, and allow to dry. Alternatively, cells can be attached to the slide by cytospin preparation. Continue with step 2 immediately below (Subheading 3.2.2.). 

Carefully follow the manufacturer s pipet cleaning protocols and... 

Transfer the pipet to a receiving vessel and drain it by gravity while holding the tip against the wall of the vessel. After the liquid stops, hold the pipet to the wall for a few more seconds to complete draining. Do not blow out the last drop. The pipet should be nearly vertical at the end of delivery. When you finish with a pipet, you should rinse it with distilled water or soak it until you are ready to clean it. Solutions should never be allowed to dry inside a pipet because removing internal deposits is very difficult. 

To clean a mercury spill, consolidate the droplets with a piece of cardboard. Then suck the mercury into a filter flask with an aspirator. A disposable Pasteur pipet attached to a hose makes a good vacuum cleaner. To remove residual mercury, sprinkle elemental zinc powder on the surface and dampen the powder with 5% aqueous H2S04 to make a paste. Mercury dissolves in the zinc. After working the paste into contaminated areas with a sponge or brush, allow the paste to dry and sweep it up. Discard the powder as contaminated mercury waste. This procedure is better than sprinkling sulfur on the spill. Sulfur coats the mercury but does not react with the bulk of the droplet [D. N. Easton, Management and Control of Hg Exposure, Am. Lab., July 1988, p. 66],... 

Contaminated glassware should be kept separated from uncontaminated. Contaminated beakers and flasks are placed in the special sink or other container for washing. Clean and wash all equipment with soap and water immediately after the experiment has been completed. If water-insoluble materials are being used, the first washing should be done with an organic solvent such as acetone. Soak contaminated pipets in a container filled with water. All broken glassware is disposed of in the Solid Radioactive Waste container. 

Standardize the spectrofluorimeter in the following way. Pipet 2.0 mL of the pH 7.5, ethidium bromide-Tris buffer into a cuvette. Add 1.0 mL of Tris buffer I and 20 fiL of standard DNA solution. Mix and place in the fluorimeter. Adjust the fluorescence intensity to 100. Clean the cuvette as described in part A and repeat the assay using various concentrations of spermine. Prepare a table displaying the amount of each component to be added. Four reagents must be in the table pH 7.5, ethidium bromide-Tris buffer, DNA solution, spermine, and Tris buffer I. Maintain the volume of DNA at 20 fiL and ethidium bromide solution at 2.0 mL for all assays. Use 0.05, 0.1, 0.2, 0.4, 0.6, 0.8, and 1.0 mL of spermine in the assays. Remember that the total volume of all constituents in the cuvette must remain constant at 3.02 mL for all the assays. Therefore, the amount of Tris buffer I must change with the amount of spermine added. Prepare each assay separately by adding the proper amount of each component to the cuvette. Mix well and record the fluorescence intensity of each cuvette. 

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