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Why electroporator need a cuvette?

Why electroporator need a cuvette?


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I have never obtain an Electroporator for my home-lab. Because it's very expensive, I'm trying to create a diy cheap version for myself. In all commercial design, I see that we need to put the sample to a 0.1cm-0.2cm cuvette to doing an electroporation, can I just replace it with an eppendorf plastic tube and put my 0.1->0.2cm alumium electrode inside?


Generally I would advise against making a home-brew electroporator: the voltage needed for electroporation is so high, that even with professional equipment you can easily get arc lightning. In an self-built device that would pose a serious fire hazard.

However even theoretically there are problems with the design you propose:

1) It is important that ALL bacterial cells in the cuvette get exposed to electrical field induced by the high voltage spike. If you use simple electrodes you will only affect the few cells directly between them, which will drastically reduce efficiency of the electroporation. For your design to work properly you would need electrodes that exactly match the dimensions of an eppendorf tube to make sure all bacteria will be squeezed between them.

2) With your device you can only do one electroporation, then you have to thoroughly clean the electrodes. This has to be done for two reasons: first you don't want any carry over bacteria (or as Chris pointed out DNA) from one batch to another. Secondly the electrodes need to be clean of any salts (or other molecules that increase conduction in a solution) to prevent arc lightning between them.


ElectroPen: An ultra-low–cost, electricity-free, portable electroporator

Affiliations School of Chemical and Biomolecular Engineering, Georgia Institute of Technology, Atlanta, Georgia, United States of America, Lambert High School, Suwanee, Georgia, United States of America

Affiliation School of Chemical and Biomolecular Engineering, Georgia Institute of Technology, Atlanta, Georgia, United States of America

Affiliation School of Chemical and Biomolecular Engineering, Georgia Institute of Technology, Atlanta, Georgia, United States of America

Affiliation School of Chemical and Biomolecular Engineering, Georgia Institute of Technology, Atlanta, Georgia, United States of America

Affiliation Lambert High School, Suwanee, Georgia, United States of America

Affiliation School of Chemical and Biomolecular Engineering, Georgia Institute of Technology, Atlanta, Georgia, United States of America


SUMMARY

The labeling of specific molecules and their artificial control in living cells are powerful techniques for investigating intracellular molecular dynamics. To use these techniques, molecular compounds (hereinafter described simply as ‘samples’) need to be loaded into cells. Electroporation techniques are exploited to load membrane-impermeant samples into cells. Here, we developed a new electroporator with four special characteristics. (1) Electric pulses are applied to the adherent cells directly, without removing them from the substratum. (2) Samples can be loaded into the adherent cells while observing them on the stage of an inverted microscope. (3) Only 2 μl of sample solution is sufficient. (4) The device is very easy to use, as the cuvette, which is connected to the tip of a commercially available auto-pipette, is manipulated by hand. Using our device, we loaded a fluorescent probe of actin filaments, Alexa Fluor 546 phalloidin, into migrating keratocytes. The level of this probe in the cells could be easily adjusted by changing its concentration in the electroporation medium. Samples could be loaded into keratocytes, neutrophil-like HL-60 cells and Dictyostelium cells on a coverslip, and keratocytes on an elastic silicone substratum. The new device should be useful for a wide range of adherent cells and allow electroporation for cells on various types of the substrata.


Electroporation cuvettes-single use only? - (Sep/23/2006 )

hi,
can electroporation cuvettes be used just once really?
i mean do you think i can use them again and again for the transformation of same bacterial line with the same DNA?

They say it is only one time use but i think it can be used juz once more if it is same DNA and the same bacterial line.
the marketing executive who came to demonstrate the electroporator said if doing the same DNA and bacterial line the same cuvette can be used again for electroporation on the same day.
gud luck. try and c

actually those single use eletroporation cuvettes can be use again, and again and again and again.

Until it cracks from miss use and rough handling.

After a cuvette is used, simply throw it into a beaker of dilute Trigen (non ionic detergent)

Once beaker is full rinse the cuvettes rigorously at least 5 times under running tap water, followed by at least 4 times with distilled water. Soak in very hot distilled water for 15mins (optional). Then shake vigourously with 70% EtOH for 20mins (we use a shaker to do the shaking). Replace the solution with fresh 70% EtOH, shake some more(20min). Do this for a final third cycle. Then drain EtOH, cover and leave in on bench for a few days for EtOH to evaporate.

we used to use he cuvettes till they cracked.

We would wash them and throw the cuvettes in beaker with 70% ethanol. We would then wash it vigorously with water for a couple of times and then finally wash it in distilled water and then again dry it.

we pretty much follow the same as mentioned above in our lab, except we expose the cuvettes to UV light (about 5 min) also.

ok here really simple, they just wash with ethanol three times and resuse directly. as everyone here is convinced to have great results, i keep reporting contamination. i am wondering if it's from the cuvettes however we only have 5 cuvettes in the lab which are constantly reused for many years so i can't try with new ones to double check, ordering of which is out of question obviuosly. anyway i am beginning to suspect that others also have contamination but dont report..

Contamination.. oh dear. It does happen with reusing cuvettes but it is usually very small.

Cuvettes can be rather cheap if you were to shop around. I am not sure how much they cost in Japan, but in UK the price can vary from 11GBP per cuvette all the way down to 1GBP. The cuvettes all look exactly the same, work the same. They even have the same colour coding. I think it is the same product just priced differently to catch people and bleed 'em dry.

So perhaps it would be an idea to go shop around. The smaller companies (at least here) tend to be alittle less blood thirsty with their prices and more friendlier/prompt with their service.

we used to get it for 1-2 euros a piece. u could try getting just a few pieces to try it out.

i have always reused electroporation cuvettes, and reused them. milliq water and 70% ethanol is sufficient as a wash. i generally squirt first with water, then with the ethanol, then again with water, dry and keep for the next round.

i do like viv, and every two month, i put all cuvettes in a bath of 0.5N NaOH for 1h to overnight and wash them then with water.
No contamination so far.
I throw them away when electric arc occurs twice with the same cuvette.


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The process

Electroporation is done with electroporators, appliances which create the electric current and send it through the cell solution (typically bacteria, but other cell types are sometimes used, as discussed above). The solution is pipetted into a glass or plastic cuvette which has two aluminum electrodes on its sides.

For instance, for bacterial electroporation, a suspension of around 50 microliters is usually used. Prior to electroporation it is mixed with the plasmid to be transformed. The mixture is pipetted into the cuvette, the voltage is set on the electroporator (240 volts is often used) and the cuvette is inserted into the electroporator. Immediately after electroporation 1 milliliter of liquid medium is added to the bacteria (in the cuvette or in an eppendorf tube), and the tube is incubated at the bacteria's optimal temperature for an hour or more and then it is spread on an agar plate.

The success of the elecroporation depends greatly on the purity of the plasmid solution, especially on its salt content. Impure solutions might cause a small explosion (known as arcing), in which case the bacteria are dead and the process needs to be redone. If this happens often, an additional precipitation may be performed prior to electroporation.


Why electroporator need a cuvette? - Biology

The gap width of cuvette I use is 0.4 cm. Can I use this cuvette for yeast and bacterial cells?

Can anyone give me the protocol of using this cuvette with bio-rad micropulser electroporator?

0.4cm gap width cuvvettes are used for mammalian cells or ES cell electroporation as per bio-rad recommendation.

did it work with with you ?

0.4 cm gap size cuvette works with E. coli.

Hi Winschau, I am interested to the transformation efficiency when the 0.4cm cuvette was used for E. Coli. Thank you! I want to try it too.

Which journal is suitable to publish ISI-low impact factor for protein modeling/bioinformatics/structural biology?

I want to publish manuscript related to protein modeling with low impact factor,indexed by ISI, scopus..

How can I predict and evaluate this DOPE profile?

Can anyone give comments for this DOPE profile? Thank you.

How to download this article? thanks.

Floral volatiles from biosynthesis to function

Hi,what is the suitable modelling server for the size of amino acid sequences, ranging from 300 to 450 amino acid residues?

I am a novice user for using protein bioinformatics tools. I don't know the clear path for analyzing my amino acid sequences of interest for modelling 3-D structure. What is the initial step of.

How can one use TSP-PRED properly for submitting protein sequences?
How to open Geno3D?
How to change to .ali file and .py file?

My current files for modeller 9.14 (Mod9.14) were all in notepad, except for protein sequences in pdb format. I dont know how to change my files to .ali and .py format. Any suggestion?

Can someone advise on purified his-tag protein in elution buffer containing 500 mM imidazole (His Spin Trap kit - GE Healthcare)?

I eluted purified his-tag protein in 200 ul of elution buffer containing 500 mM imidazole. Then, I want to use it for SDS-PAGE to confirm the presence of specific his-tag protein band on gel. What.

Heartburn. Is it an early warning sign of cancers?

Is it true that the use of H2 blocker, proton pump inhibitors and other antacids can lead to the progression of recurring heartburn? The effect of reducing stomach acid can also make gastrin to be.

Separating gel does not solidify. How to troubleshoot?

Is it necessary to put any liquid on top of TEMED-treated separating gel mixture between spacer plates with 1.0 mm Integrated spacers in closed casting frame?If yes, what type of liquid.

Fatal error

When I tried to energy minimization my system, I got fatal error as below. Fatal error: Atomtype opls_116 not found Although I've already added this line: include water #include "oplsaa.ff/spc.itp" to [molecultype] directive in my topology.

Fatal error

When I tried to energy minimization my system, I got fatal error as below. Fatal error: Atomtype opls_116 not found Although I've already added this line: include water #include "oplsaa.ff/spc.itp" to [molecultype] directive in my topology.

Fatal error

When I tried to energy minimization my system, I got fatal error as below. Fatal error: Atomtype opls_116 not found Although I've already added this line: include water #include "oplsaa.ff/spc.itp" to [molecultype] directive in my topology.

Fatal error

When I tried to energy minimization my system, I got fatal error as below. Fatal error: Atomtype opls_116 not found Although I've already added this line: include water #include "oplsaa.ff/spc.itp" to [molecultype] directive in my topology.

What kind of fluid could I use for a pitfall trap that also does not invalidate molecular testing?

Hi, I want to start testing pitfall trap to obtain ants samples, but I need to conduct molecular analysis on those insects. So, what kind of fluid can I use? Ethanol expires too early and I need.


Part I: Identifying Food Dyes in Candies

Many foods, drugs and cosmetics are artificially colored with federally approved food dyes (FD & C dyes). These dyes include Red 40, Red 3, Yellow 5, Yellow 6, Blue 1, and Blue 2. Since each dye has an identifiable absorption spectrum and peak, a spectrophotometer may be used to identify the types of FD & C dye used in a product.

Pigments may be extracted from foods and drinks that contain one or more of these dyes. An absorption spectrum of that extract can then determine what dyes are in that food or drink by comparing the peaks of maximum absorbance with information in the table below. If the absorption spectrum of a food extract has a peak at 630 nm and one at 428 nm, you can assume the food contains both Blue #1 and Yellow #5. The following table gives the wavelength of peak absorbance for each of these dyes.

Table 1. Wavelength of Maximum Absorbance of Commonly Used FD & C Dyes

Materials

Procedure

A. Extracting Dye from Candy (Your Instructor will do this for you)

You will need one test tube and one cuvette for each color to be tested. Measure 4 mL water into one tube. Place 2-4 candies of the same color in a test tube with the water. Gently swirl, and wait one minute. After, pour approximately 1 mL of liquid into a microcentrifuge tube. Spin the microcentrifuge tube at max speed for 60 seconds. Make sure the centrifuge is balanced before spinning. Transfer the clear liquid (supernatant) into a cuvette. Make sure to leave behind the particulates (pellet).

Figure 4. Curve of absorbance


A novel electroporation method using a capillary and wire-type electrode

Electroporation is widely used to achieve gene transfection. A common problem in electroporation is that it has a lower viability than any other transfection method. In this study, we developed a novel electroporation device using a capillary tip and a pipette that was effective on a wide range of mammalian cells, including cell lines, primary cells, and stem cells. The capillary electroporation system considerably reduced cell death during electroporation because of its wire-type electrode, which has a small surface area. The experimental results also indicated that the cell viability was dependent on the change in pH induced by electrolysis during electroporation. Additionally, the use of a long and narrow capillary tube combined with simple pipetting shortened the overall time of the electroporation process by up to 15 min, even under different conditions with 24 samples. These results were supported by comparison with a conventional electroporation system. The transfection rate and the cell viability were enhanced by the use of the capillary system, which had a high transfection rate of more than 80% in general cell lines such as HeLa and COS-7, and more than 50% in hard-to-transfect cells such as stem or primary cells. The viability was approximately 70–80% in all cell types used in this study.


Transfection vs Electroporation

Whether using viral or nonviral reagent-based methods of transfection, or the more modern instrument-based methods such as electroporation, in theory the results should be more or less similar. Regardless of the path we take to get there, the destination should be the same. However, as we all know, that's not always the case. Both methods can produce results. Why? And which method is best?

In terms of flexibility, both methods allow for a little variation in process. Both enable different applications to be utilized to obtain the desired result ultimately, but we are starting to see a definite trend for specific applications in particular. For transfection, for example, nonviral vectors like lipoplexes are believed to be more successful and have lower risk factors than viral vectors. And in electroporation, single cell electroporation processes (SCEP) are typically favored over bulk electroporation processes (BEP) due to higher cell viability, greater transfection efficiency, and a lower rate of contamination.

Comparison of Methods

How well does each method work? When it comes to transfection as a whole, cell viability and cell toxicity are primary concerns. Both processes are at risk for reduced cell viability. Transfection, most notably, due to an incorrect transfection reagent ratio, the use of cells with lower densities of 70 percent or less, and inadequate transfection reagent storage (which should be 39.2 degrees). Cell viability in electroporation is largely dependent on waveforms, field strength, cuvette gap size, cell diameter, temperature, pulse length, and the number of pulses, but may be influenced by many more factors.

The primary difference in risk factor appears when looking at cell toxicity. One of the most prominent and well publicized advantages of using an electroporator is, of course, that these instrument-based processes manage to avoid many of the main advantages of viral or nonviral reagent-based transfection, including cell toxicity. Unfortunately, cell toxicity is still an important factor to consider in reagent-based transfection. Cell toxicity can significantly affect results, and some reagents, particularly liposomal transfection agents, are well known for activating stress response pathways, affecting cell regulation.

While both methods allow for both DNA and mRNA transfer, there are advantages and disadvantages of both techniques that should always be taken into account. A main advantage of electroporation, for example, is that it boasts high levels of reproducibility, while replicating findings in reagent-based transfection can be challenging, due to splits and changes in cells. On the other hand, reagent-based transfection is a much more cost effective solution, especially when calcium phosphate or cationic polymers are used. The time differences for cell harvest between the two processes are negligible.

Which transfection method is superior? It's a difficult question to answer. However, it's no secret that many are now opting for electroporation over reagent-based transfection, because it lacks the same level of restrictions that reagent-based transfection has, most notably in terms of the type of cells that can be targeted. We're also seeing a shift due to the limitations in length of the DNA that is inserted with reagent-based transfection, and because of emerging issues with biosafety. However, that's not to say that reagent-based transfection is dead. Far from it. In fact, one of the more recent trends in transfection is to hybridize techniques to increase efficiency and viability, and reduce toxicity and other risks.

At VitaScientic, we can provide solutions -- no matter the option you decide is best for you.
To learn more about the new and innovative Celetrix sealed-tube Electroporation System: Click here >>
To see a complete list of the best-in-classs DNA-In&trade and mRNA-In&trade Transfection Reagents: Click here >>


Watch the video: Proper placement of cuvette in a Beckman spectrophotometer (July 2022).


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