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Hey everyone. I am trying to find a cheap mix of lipids to use as a system suitability standard to make sure the instrument and chromatography is working as expected. What do people use? And most importantly, how much (volume/concentration/amount) do people load for different sized columns and flow rates?
Thanks.
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by fractional inhibitory index
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We are planning to synthesize customized liposomes for the purpose of delivering nucleic acids into bacterial cells.
The methodology requires us to prepare a thin film of lipids (Cardiolipin and DSPC). For this step, we have to take the above lipids in a cholroform solution and evaporate the solvent using an rotary evaporator followed by hydration with HEPES buffer and sonication to yield liposomes.
However, we face challenges in setting this reaction in a conventional rotary evaporator. The round bottom flask thats routinely used with the rotary evapotor is 500 ml. Our reaction volume is around 1-2 ml only. Also I'm unsure whether I would be able to perform the sonication step with the bigger round bottomed flask.
It would be easier if i can setup this reaction in an eppendorf tube using a Vaccum centrifugal concentrator. For this reason, I would like to know whether the lipid thin film synthesis protocol could be executed in an ependorf tube.
Someone having expertise in liposome synthesis and delivery kindly help me.
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I would recommend using a single-neck round-bottom flask suitable for smaller volume like https://www.sigmaaldrich.com/NL/en/product/aldrich/z723134
The thing is that because of the use of chloroform the combination with plastic should be avoided as much as possible. See for examples how resistant (or not) polypropene is: https://www.engineeringtoolbox.com/polypropylene-pp-chemical-resistance-d_435.html
So, there is too much risk that an organic solvent like chloroform will dissolve (or better put extract) all types of unwanted substances like organic plasticisers, phthalates, BHT. Using glass is a much safer way to avoid such unwanted side-effects.
Best regards.
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I am making LNP, and I need to track my particles after in vivo injection. So i need to lable the LNP with a stain like Dil stain. However, I don’t know how I am going to incorporate the stain in the LNP, or what is the technique to do that ?Also, I need to know how to check if the stain successfully incorporated into the particles. Does any one has experience in using this Dil stain with liposome or lipid based particles?
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Hi Aziza, I have this same question and was wondering if you were able to incorporate it in your LNPs and how did you check for incorporation? Many thanks!
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Hello everyone,
My lab plans to use Phospholipid Fatty Acid Stable Isotope Probing (PLFA-SIP) to see if a specific microbe utilises a certain substrate as a carbon source. Luckily our microbe has a unique lipid molecule only found within its group.
However, we are trying to figure out what analytical machine will be best for our purposes. I have found in some literature using gas chromatography/isotope ratio mass spectrometry (GC-c-IRMS). Therefore, would this be suitable for detecting potential isotope uptake in our microbe?
Thank you very much
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Hello Dr. Noel Davies,
Fantastic! Thank you very much for telling me this and for your help it is much appreciated!
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I want to prepare lipid nanoparticles? Are there any protocols or papers about using ultrasound?
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Dear Yue Zheng, please have a look at the attached files. My Regards
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The presence of C12 acyl chain makes it difficult to separate it through the conventional molecular sieve chromatography!
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Hi Rashmi,
You can try fractional separation with no-polar solvents such as pet. ether and further purify using semiprep HPLC.
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I'm doing invitro drug release using the dissolution test for betamethasone dipropionate lipid nano particles. I used PBS PH 5.5 with 1% tween 80 ( 37C heat) according to previous articles. However the betamethasone dipropionate was not soluble. All drug was still entrapped in the dialysis bag. I also tried dissolving betamethasone dipropionate of same concentration as in the lipid Nano particles ( free powder form) in PBS PH 5.5 with 1% tween 80 ( 37C heat) and still didn't dissolve.
I also tried reducing the PH below 5.5 and increased the tween % but still the same. I used PBS: Ethanol 70:30 and still didn't work noting that alcohol may break the lipid nano particles.
Any suggestion for a solvent to dissolve the betamethasone with out disrupting or affecting the lipid nanoparticles??
Thanks,
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You are most welcome dear Tia Mo . Wish you the best always.
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I was wondering if anyone had any experience with measuring lipid (fatty acids) using tissue that has been stored in RNAlater?
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I worked with placenta tissue stored in RNAlater. This storing method is not a problem for F¡fatty acid extraction neither analysis with GC.
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Hi everyone, I am using an inverted emulsion method to generate my liposome (POPC), but there were a huge number of non-liposome clumps under the microscope. I don't know why this can happen and in which step I was wrong. So I am asking how can I get rid of these clumps?
The phospholipids I am using is POPC and Texas Red as dye.
Also, I used beta-cacein as surface coating agent.
Centrifugation parameters were 400g 10min.
My protocol was as below:
1. Preparation of lipids in oil solution: POPC + Texas Red + mineral oil, with the final POPC concentration of 200 uM
2. Prepare the inner aqueous solution: 600mM sucrose
3. Prepare the outer aqueous solution: 600mM glucose
4. Prepare interfacial lipid monolayer: 50 μL of glucose (outer) solution were added to the pre-coated wells in the microtiter plate. Add of 20 μL of the lipid–oil mixture on top of the glucose. The entire setup was allowed to incubate for a period of 30 min to form an interfacial lipid monolayer
5. Prepare the water in oil emulsion: 250 μL of the same lipid–oil mixture were added to a 1.5 mL Eppendorf tube. To this tube, 10 inner solution was added and then agitated mechanically to yield a water-in-oil emulsion.
6. Prepare the GUVs: An aliquot of 50 μL of the emulsion was pipetted into the wells containing the lipid monolay- er interface. Immediately after this, the microtiter plate was transferred into a centrifuge (400g 10min)
Thank you!
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Please explain in detail what you want, I can't understand what you're asking!? We have technology and chemists to deal with the formation of liposomes, and I am a pharmacologist!
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Hi
I want to MD SLN in gromacs first of all i have putted my lipid molecules in a box and then added drug molecules. Now I want to energy minimize; for this work I need a topology file but I don't know how should I create it.
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Where do you want to create the file? In this platform?
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Hi everyone!
I am trying to encapsulate a metallic photosensitizer along with 5-Fluorouracil (5-FU) inside a liposome. The preferable size is 30 nm.
In brief, the preparation of liposomes is carried out by the thin-film hydration method where the drugs and the vesicles are incubated above the phase transition temperature of lipid. After that, to downsize the liposomes, several freeze-thaw cycles are performed before sonication followed by filtering it with a 0.03 µM PCTE membrane filter.
However, it seems that our liposomes are leaky, as there is no difference in 5-FU content between the filtrate and the liposome, as indicated by HPLC. Additionally, during the washing of unentrapped molecules with 5% glucose solution I observed some debris on the wall of the filtering unit, most likely due to the lysis of liposomes.
Furthermore, I also need a suggestion regarding the suitable size of liposomes to avoid liver and spleen toxicities.
I have no expertise in the field of liposomal formulation. Therefore, any input or clarification would be highly appreciated.
Thank you!
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Jiun Wen Guo, thank you for your suggestion.
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I wish to create unilamellar liposomes of ~0.2 microns and have seen mini-extruder options such as the Avanti Lipids Mini-Extruder set (https://avantilipids.com/divisions/equipment-products). My lab has syringes and syringe filter units of comparable pore size (0.22 microns) capable of up to 145 psi inlet pressure (https://www.emdmillipore.com/US/en/product/Millex-GP-Syringe-Filter-Unit-0.22m-polyethersulfone-33mm-gamma-sterilized,MM_NF-SLGP033RS#anchor_keySpecTable).
Taking cost into consideration, is it acceptable to use the syringe+filter apparatus as an alternative to this mini-extruder? If so, what special precautions would I need to take, if any?
Side note, I will not need to heat up the suspension (i.e., room temperature is fine). Any help or new ideas are appreciated!
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It can be used if your vesicles are "fluid" enough. Do you have any sterol in your formulation?
The other parameter is to make sure vesicles are passed through filters at temperatures above their Tc.
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Dear all,
I would like to perform a solid phase extraction to obtain a lipid extract from dried blood spots. For that reason, I would like to know what particle size of silica gel should I buy? This lipid extract would be analyse by LC-MS/MS.
Thanks you all,
Best regards.
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if You are going for (offline) SPE I do agree to Elias. You could also imagine running a real separation column (online) than You would rather work with 4 µm particles.
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in downstream processing of biopolymer some quantity of protein still remains even after treating enzymatically which are known for protein removal.
looking for complete removal of protein from polymer, please suggest commercially feasible options.
protein source is bacteria whose outer layer consist of lipids and protein
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Dear all, the conventional way is via 'sevag method for deproteinization'. Please make simple google search using the keyword 'methods of deproteinization'. The following sample documents may help. My Regards
10.4028/www.scientific.net/AMR.340.416
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I am working with yeast lipids. After LCMS-IT-TOF and MS-dial analysis I got the list of lipids. i need to know which of these lipids belong to yeast. I was looking for a database that can help determine list of yeast lipids. I found yeast metabolome database , I guess that might not be the appropriate one.
Thank you.
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Thank you. It was helpful
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I want to optimize the synthesis of nanoparticles using DOE.
My system is: lipid A and lipid B (total mass of two lipids stays the same), surfactant and water.
Which DOE should I choose?
  • Mixture - to make two lipids as a Mixture A and surfactant and water as a Mixture B?
  • Combined - to make two lipids as a Mixture A and surfactant as a numeric factor (not mentioning water here)
Just to be clear. Total mass of lipids does not change, only the ratio. Then I want to check different masses of surfactant (w/w surfactant/total lipids mass). Water is added, so total mass of sample is constant.
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This was my first idea, however I was concerned that such approach will interfere with calculations.
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membrane lipids
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It depends on what you want exactly but I guess a good starting point might be: L J Macala, R K Yu, S Ando, Analysis of brain lipids by high performance thin-layer chromatography and densitometry., Journal of Lipid Research, Volume 24, Issue 9,
1983, Pages 1243-1250, ISSN 0022-2275, https://doi.org/10.1016/S0022-2275(20)37906-2. It is an open access paper (with some good references to other relevant papers).
Best regards.
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Currently, liposomes are synthesized using lipid-coated beads. To make lipid-coated beads, lipid powder is initially dissolved in methanol containing rhamnose and glass beads. I am wondering about the function of the rhamnose, is it related to the coating ability of lipid on the beads?
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Kindly see also the following useful RG link:
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We made a pilot study with N=2 mice per group to test biodistribution of rhodamine 6G-loaded silica nanoparticles functionalized with PEG, at 1.5h, 6h and 24h. We used GFP detector in IVIS Spectrum 200 imaging system and normalized fluorescence efficiency to control organs average, as a ratio (N=2 as well).
We expected to find high accumulation in liver, spleen and lung, possibly decreasing with time, but surprisingly we cannot see any rhodamine signal in liver at any time when compared to control organ. We made sure the biliary vesicle was not present and liver was properly washed. Signal in spleen is not very high either. Nevertheless, lung signal at 1.5 hours is 5-fold the controls signal, decreasing in time and disappearing at 24h.
We know optical imaging is not the most sensitive technique to assess nanoparticle biodistribution, but we can clearly see rhodamine signal in lung. What could possibly be going on in liver and spleen? Could their color or opacity cause any quenching of the signal? I've found this article detecting rhodamine in liver using the same equipment as we do () so I guess it is not a matter of quenching...
Do we have the best and most biocompatible nanoparticles ever? I don't think so! But we need an explanation for the lack of liver uptake...
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Dear Rob Keller,
Thank you very much for your kind answer!, I did not realise there was such a difference!
Still, to avoid undesired interactions that could lead to different biodistributions (nanoparticles like ours are usually uptaken by RES organs), or dye degradation, in our case rhodamine is loaded *inside* the nanoparticle. Thus, it is not exposed to plasma, so I do not believe it is a matter of solubility (at least, not rhodamine-related)...
I'll keep on researching!
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I am trying to bind lipid extractions as well as purified single lipids from mycobacteria to Nunc PolySorp 96 well plates to perform ELISA's.
Lipids are insoluble in water, so when I try and dilute down my lipid fractions (in 2:1 chloroform:methanol) into carbonate buffer I am seeing them precipitate out.
I have tried air-drying the lipids at RT after adding to the plates in 2:1 chloroform: methanol as well as pure methanol but I haven't had much luck.
Does anyone have any suggestions or experience on how to bind lipids to ELISA plates?
Thanks,
Nathan
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I agree with my colleaques
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Hi Everyone
I am trying to calculate needed lipid amount from initial D:L ratio prior to remote loading.
I have a drug compound with a concentration of 1mg/mL (volume=1ml) with an MW=521.6 which i am going to load into liposomes.
I have an initial 0.5mM lipid stock solution.
I have calculated the compound concentration to be 1917micromolar which gives me for a Drug-to-lipid ratio of 1:4 an addition of 15.3mL from the 0.5mM lipid stock.
However it seems quite high amount of lipid and i am therefore not sure of the calculations are correct.
Any input or clarification is highly appreciated
Thanks!
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Hi,
This seems about right. You initially have 1 mL of a 1 mg/mL solution of the drug, which has a MW = 521.6 g/mol, so you have indeed a 1916 uM or 1.916 mM solution of the drug.
When you add the 15.3 mL of the 0.5 mM lipid solution to your 1 mL drug solution, you end up diluting the drug solution to a concentration of 117.62 uM (1917 uM/(1 mL + 16.3 mL) = 117.62 uM).
Your initial lipid solution (15.3 mL) has a concentration of 0.5 mM, but you slightly dilute this solution with the 1 mL of drug solution, you end up having a concentration of 0.46933 mM or 469.33 uM of the lipid (0.5 mM * (15.3 mL/16.3 mL) = 0.46933 mM).
Since you have 117.62 uM of the drug and 469.33 uM of the lipid, you have 117.62 uM drug/469.33 uM lipid = 0.2506, which is indeed about 1/4.
It may seem like you have way too much lipid, but this ratio makes sense. You start with about 2 mM drug solution and a 0.5 lipid solution, so you have about a 4:1 drug : lipid concentration initially, but you have 15.3 mL of the lipid solution and only a 1 mL drug solution, so you have about a 1:16 drug : lipid volume. When you take both a volume and the initial concentration into account, you indeed have a 1:4 drug : lipid ratio.
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I am analyzing a batch of adipocyte tissues using Adiposoft which is a plugin for Fiji. The program is taking the area of the cells and giving me their Equivalent Diameters (D-eq). I read that "From the diameter, the average adipocyte volume and lipid content can be mathematically derived" (Galarraga et al. 2012).
Could someone tell me what formulas (and the definition of their variables) I can use to find the Lipid content and the volume of an Adipocyte using the D-eq and Area?
I also have the size of the image in microns and an estimate of how many cells are in the image based on what Adiposoft counted during its automated analysis.
Galarraga, M., Campión, J., Muñoz-Barrutia, A., Boqué, N., Moreno, H., Martínez, J. A., Milagro, F., & Ortiz-de-Solórzano, C. (2012). Adiposoft: automated software for the analysis of white adipose tissue cellularity in histological sections. Journal of lipid research, 53(12), 2791–2796. https://doi.org/10.1194/jlr.D023788
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Wait since its the equivalent diameter do I just calculate the equivalent volume using the volume of a sphere formula? -_-
V-eq=(1/6)*Pi*(D-eq)^3?
And is the equivalent volume then equal to the lipid content?
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I want to learn what is the reaction factor or constant for liposome formation. We are going to simulate it by COMSOL, Diluted Species Model. But we need a factor for the addition of lipids. Maybe I asked this question wrong, but it will be very helpful if you answer. Thanks!
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Good luck
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i have a nano particle and lipid system and i want to move the nano toward the COM of lipid. how can i perform this in GROMACS? the motion is in the Z-axis
it will be preferable if there is an mdp file or a link
thanks
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I was just wondering this when is was reading about media like Mackonkey. Just out of curiosity, I read few of the articles.
Is it due the presence of teichoic acid in the CW of gram positive organism that contains lipid? or something else, protein aggregation? How is the selectivity obtained?
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I was also wondering about this! We use MacConkey agar to kill gram positives/select for gram negatives, but if bile disrupts lipids it would seem that gram negatives would be more vulnerable via their outer membrane. I would like to hear from true experts, but I did find this article useful-- it does seem that bile exerts its antimicrobial effects through proteins, not lipids:
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How does the temperature affect the shape of the lipid, if we have, for example, a phospholipid with unsaturated chains, will the shape of this lipid change at a higher temperature? Is there a greater chance of getting e.g. reverse micelle. I do not know if this is true, but I heard somewhere that the higher the temperature, the movement of the unsaturated acyl chains increases, indirectly increasing the volume of the non-polar part of the lipid. Is it true? Do you know any literature references on this topic.
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Dear Jakub Hryc ,
The whole concept of lipid shape and lipid polymorphism basically started in the following classical paper: Cullis, P. T., & De Kruijff, B. (1979). Lipid polymorphism and the functional roles of lipids in biological membranes. Biochimica et Biophysica Acta (BBA)-Reviews on Biomembranes, 559(4), 399-420 (see enclosed file, especially Figure 7).
Another excellent (more recent) paper is: Frolov, V. A., Shnyrova, A. V., & Zimmerberg, J. (2011). Lipid polymorphisms and membrane shape. Cold Spring Harbor perspectives in biology, 3(11), a004747. https://cshperspectives.cshlp.org/content/3/11/a004747.short(scroll down a little bit and you find an impressive collection of related papers written by the ‘big’ names in the field).
Best regards.
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We have done ELISA quantitation of a serum protein in a total of 200 patient samples across five different sample categories. We had assessed the blood parameters such as ESR, CRP, lipid profile for all these patients.
For the ELISA data, I did a Kruskal-Wallis non-parametric test to check if there is any significant difference between the groups. I also did Dunn's test for pairwise comparison to identify which groups were statistically significant.
For one particular protein, the comparison of the protein values did not show any statistically significant difference across any pair of groups. However, when I normalized the protein concentration with the ESR for each patient and compared, the protein/ESR ratio showed statistically significant difference between some pairs of sample groups.
Is this kind of normalization acceptable for doing statistical analysis?
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Dear Jeya,
I think you better normalize your data with the total protein concentration of your samples. ESR and CRP are factors that change relatively much in different persons and their levels depends on many factors. Thus, they cannot be used as reference factors. Personally, I set all my samples at the same protein concentration for an ELISA test. I also draw a standard curve with the concentrations of my standard samples. This ensures you that you have a low variability.
Good luck
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Dear all,
I want to exposure my human primary cell line (fibroblasts) to lipids. There is not much literature available outlining how best to do this. Has anyone experience of doing so?
Specifically do I need to prepare a liposome first and if so, what do I use as the carrier/ buffer?
Thanks in advance,
Marissa
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I'm sorry I don't have direct experience with that. Good luck!
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Hello.
I'm a starter for biomass and biodiesel. I have a question about biodiesel.
For making biodiesel, we are reacting lipids such as animal fat, soybean oil, or some other vegetable oil with an alcohol to produce a methyl, ethyl, or propyl ester by the process of transesterification,
Here, I have questions.
1. Why do we have to make the ester molecules for biodiesel? In that, we can't use lipids as it is for biodiesel?
2. What is the function of the ester in biodiesel?
I will wait for your reply.
Thank you so much.
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1. due to the fact that ester molecules have a much lower viscosity than triglyceride molecules due to its smaller molecular weight. Triglycerides, on the other hand, can be used directly as a biodiesel. Due to its high viscosity, the issue is that it will eventually cause damage to the fuel injecting system.
2. The importance of the ester is that it converts a high-viscosity triglyceride to a low-viscosity triglyceride that may be used as biodiesel without any problems.
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Hello,
I need to perform MD simulation for a protein embedded in a mixed membrane bilayer with certain concentrations of different lipids (sqd, dgd, and mge lipids).
I didn't find tutorials for how to construct the lipid bilayers from scratch for MD simulation using the abovementioned lipids, anyone can help?
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Thanks Stéphane Abel for your Answer, I build the bilayers using packmol. It is actually taking time and not straight forward method :D, so I will try your suggestion it might be helpful later :)
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I wish to study the effect of autophagy inhibition on lipid accumulation in HepG2 cells when treated with free fatty acids (palmitic acid and oleic acid). I have observed that HepG2 cells when treated with free fatty acids have increased levels of lipid accumulation. My treatment compound reduces this lipid accumulation. It also induces autophagy.
However, when I try to inhibit autophagy with chloroquine and 3-MA, lipid accumulation reduces even further. Inhibition of autophagy significantly reduces lipid accumulation in my experiments. Whereas, available literature suggests inhibition and impairment in autophagy leads to an increase in lipid accumulation.
Can someone guide me through this discrepancy in data? Looking forward to your kind inputs. Thank you in advance.
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I think it is quite common in adipocytes. Inhibition of autophagy using BafA1, CQ or similar at early days of maturation seems to have a strong effect on trout adipocyte differentiation "blocking" both, phenotype changes and gene expression. It has also been reported in mammals.
I would assume something similar may occur in hepatocytes. I hope this helps :)
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Now LNP is the most famous star over the world thanks to ionizable lipids, like ALC-0315, SM-102 and MC3 (Onpattro, approved by FDA in 2018), which are neutral at physiological pH and trun to positve charge at a low pH. Just like other pH-sensitive materials. Although pH-sensitive liposomes encapsulating RNA have been developed for decades.
So, why these ionizable lipids and not other pH-sensitive materials? And why did it take so long to apply in clinic.
emmm....It's hard to describe my confusion excatly....
Just like a sad feeling about active targeting nanoparticles, which also have been developed for many years and have no application in clinic yet.
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Sadly, the Pfizer-BioNTech LNPs use cationic lipids, not ionizable lipids. It's quite shameful given the twenty-plus years of knowledge about the toxicity.
Unfortunately, the manufacturers of the LNP-based COVID vaccines go out of their way to avoid explaining the reality of their formulations. It took me quite a lot of detective work looking at papers and patents to find the truth.
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I have two treatment groups with 4 biological replicates each. I measured 100 lipid species in each of them and want to visualize the differences using a volcano plot.
Which of these two ways is the correct to process my data:
  • Calculate for each lipid species the averages, fold changes, adjusted p-values between the two treatment groups and then at the end log2 transform the fold changes and -log10 transform the p-values for plotting
  • Log2 transform all the measured lipid concentrations first, calculate the averages, fold changes, adjusted p-values between the two treatment groups. Plot the fold changes without further transformation and -log10 transform the p-values for plotting
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The latter is preferable to the former.
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Currently I'm trying to adjust Oil Red O staining method for liver lipids. Base protocol is A. Mehlem's (Nature Protocols; Vol. 8, No. 6; 2013; p. 1149-1154). I tried different times of ORO staining and counterstaining with hematoxylin. Had some problems with non-specific staining of cracks between cells which might have been solved with varying times. As I believe in the pics the darker dots are lipids and red colour between cells is other type of lipids? Maybe someone who has experience with ORO staining method could explain if attached pictures looks good and if not maybe suggest some corrections which might improve staining?
Liver samples are from chickens treated with high concentration of TBT.
Sample 7: 5 min. in ORO, 15 s. HTX, 30 min. wash; S8: 5 min. ORO, 1 m. HTX, 60 min. wash; S12: 5 min. ORO, 3 min. HTX, 30 min wash; S13: 5 min. ORO, 1 min. HTX, 30 min. wash.
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Reagents need to be filtered (Oil Red O).
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-Oil fortification with a hydrophilic extract
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Thank you again dear respectable colleage.
Best regards!
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I'm working on the formation of hybrid vescicles (LUVs) formed by lipids and amphiphilic block copolymers.
These hybrid membranes can either be well mixed, giving homogeneous properties across the surface of the vesicles or phase separated into lipid-rich and polymer-rich domains, which give rise to textured vesicle morphologies (aggregative distribution).
So my question is: how can I monitor the surface of LUVs by evaluating the distribution of polymers/lipids and see If I’m faced with a well-homogeneous or "aggregate" system in polymeric and lipid domains?
Since they are not GUVs, fluorescence microscopy should be excluded, so which techniques can be used?
Thanks in advance.
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Dear Caterina Presutti, in similar cases, SEM is the best techniques to visualize the state of phases repartition. Sometimes staining is even more important to define the contour boundaries of the different phases. My Regards
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I am doing lipid research from extracellular vesicles, and in near future I am going to use Mass-Spec. I am still planning my experiment. I read some literatures but I found it is little confused for me to figure out which solvent should I use to dissolve my lipid samples. Generally, I after I extracted lipids from my samples using mixture of chloroform and methanol, I will need to evaporate the organic solvent through speedvac/nitrogen evaporator to prevent the lipid from being oxidized. And if my column is Thermo Accucore C30 column, my mobile phase is A:(60% acetonitrile: 40% H2O with 10 mM ammonium formate and 0.1% formic acid) and B: (90% isopropanol: 10% acetonitrile with 10 mM ammonium formate and 0.1% formic acid). Is there any suggestions for me to try about the solvent to dissolve the lipid in before inject into Mass-Spec? Thank you ahead!
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Dear Yiwen,
Degradation of your sample by lipid oxidation is something to (always!) bear in mind when analysing lipids as this (unwanted) reaction is likely to occur during lipid extraction, storage, and analysis. The best option is to minimise it... I normally add BHT (50ug/mL) to MeOH used in the extraction.
Regarding the solvent to dissolve your extract before LC-MS injection you should remember that, as you say, you used MeOH and CHCl3 for the extraction so the lipids in your extract are soluble in those solvents. So, I would just dissolve it in a MeOH (which has similar polarity to AcN). Lipids by its characteristics tend to be apolar and thus not soluble in water
Hope this helps.
Ana
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During DNA extraction, cells are lysed and unwanted biomolecules (proteins and lipids) are removed to obtain DNA. But what happens to the RNA in this whole process? how is it removed?
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Thank you for your response Nooralhuda Alfatlawi
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I'm working on development of drug loaded lipid based topical carbopol based hydrogel for future treatment on diabetic wounds. It includes the characterization study regarding the former things and looking for a journal within impact 1-2. Kindly help me suggesting suitable journals.
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Below are some suggestions within the scope and JCR you need:
Carbohydrate Research (JCR: 2.1)
Journal of Food Processing and Preservation (JCR: 2.2)
Protein & Peptide Letters (JCR: 1.9)
Current Nanoscience (JCR: 1.8)
Medicinal Chemistry Research (JCR: 1.9)
Applied Biological Chemistry (JCR: 1.8)
International Journal of Peptide Research and Therapeutics (JCR: 1.9)
However, I suggest you access the following pages below. They are specific tools of the best publishers, which enable authors to find appropriate Journals for their submission. It's easy to use, briefly, you add the title and abstract of the paper and some Journals that publish works within the scope of your work will be suggested.
I hope this helped you.
Journals Elsevier Finder
Wiley Online
Taylor & Francis
Springer
Regards,
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Does anyone know how to calculate the molar concentration of a mixture of lipid, such as soybean polar extract?
I am doing nanodics with soybean polar extract. as you know, the ratio of membrane proteins, MSP and the lipid is much important. soybean polar extract is a mixture, how can we quantify the molar concentration? if you know, please. thank you very much for your help.
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Dear colleague,
I am testing on formulation of cationic lipid nanoparticles. The lipid compostiin is DOTAP: DSPC: Cholesterol: PEG2000 DSPE=50: 10: 39: 1. The buffers of 50 mM sodium acetate at pH 4 and 1XPBS, containing 0.9% NaCl, at pH 7 are used as the aqueous cargos. Could I have precious advice about how to reduce lipid nanoparticle size down to 100 nm (80-120 nm) with PDI less than 2. Thank you.
Sincerely,
Jacky
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Dear Jacky,
Do you pass your NP trough an extrusion filter under high pressure after you add the acques phase to your lipidic film? I believe this is kinda necessary to ensure you have small particles at the end. You will have to start from a 400nm filter and slowly go down (untill 100nm)....the more you pass trough the filters, the lower will be the pdi (you want it lower than 0.2, not 2). You could also try using a 0.5% of PEG rather than 1 and maybe lower the Dotap to 20-40%. Another option is sonication (which is usually done before extrusion). Another option is a centrifugation, to remove the bigger size NP formed (12000rpm for 10min should be sufficient). Then take the supernatant and check with DLS.
I hope this helps
Regards
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I want to extract certain lipids, which are prone to (per)oxidation, from mitochondria. To prevent them from oxidizing, I want to add BHT (Butylated Hydroxytoluene) to the isolation buffer after harvesting the cells.
What is the best concentration of BHT? I have found values ranging from 50 µM to 6500 µM, and I am confused as to what is the best amount. I want to dissolve the BHT in DMSO, is that good? Or would ethanol be better? Which substance has the least negative impact on LC?
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Dear Christina Oettmeier although I'm absolutely no specialist in this field, I just came across the following potentially useful literature reference which might help you in your analysis:
Development and validation of a reverse phase-liquid chromatographic method for the estimation of butylated hydroxytoluene as antioxidant in paricalcitol hard gelatin capsule formulation dosage form
Fortunately this paper is freely available as public full text on RG. Also please have a look at the attached article in which the authors use a concentration of 0.01% BHT:
A comprehensive comparison of four methods for extracting lipids from Arabidopsis tissues
I hope this helps. Good luck with your research and please stay safe and healthy!
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Hello to everyone,
We are study a recombinant protein (Escherichia coli) which has a huge hydrofobic cabity in its structure. By Electron microscopy we saw a extra density in this cavity and we decided to analyzed the content by Maldi-Tof. The result is in this cavity there are 3 differents types of molecules: Wax esters, phosphatidylcholine and sphingomyelin.
The questions are:
1) Is these molecules an usual contamination of recombinant proteins when we analyzed the samples by Maldi?
2) Which is the procedence of theses molecules? because they are present on animal cells and not in procariotes like E. coli?
We used a normal LB medium to express the protein ( nitrogenus bases, yeast extract and Sodium chloride) with E. coli DH5alpha (DE3). We extract the protein by cell press and purify in 2 steps: Ni-column and Gel filtration using as buffer: Tris 20mM pH: 7.4 and NaCl 150mM.
Many thanks!!!
David
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Have you tried expressing your recombinant protein growing E.coli in a defined medium instead of LB?
Yeast extract is a complex mixture and it might contain undesired contaminants that does not interfere normally with protein expression in E.coli but in your case it could be a potential source of artifacts in your protein ligand. Other sources can be your purifications columns, make sure to clean them properly because it might be another source of those lipids (I don't really think columns are the problem but it might be).
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Hello!
Anyone has experience with lipidTox staining of hepatocytes that are treated with lipid and subsequently mounted onto slides ?
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Hi! I dont have specific experience with hepatocytes, but have done some lipidtox staining. Could you describe in more detail what you have done and what is the problem?
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Hi all!
I need some help, as I've been trying to troubleshoot and struggling with this for more than a few months.
I have DOPC + 05% Texas-Red-DHPE liposomes that I need to fuse into planar lipid bilayers, but the fusion doesn't happen once they are applied on my glass slides.
My liposomes are produced using the sonication, freeze-thawing and extrusion procedure. I have confirmed their size using Dynamic Light Scattering and they are < 100nm.
My supported bilayer formation were checked using FRAP and QCM-D, both indicating that there is liposome adhesion but NO fusion.
Additional relevant info:
- Lipids are rehydrated in PBS with and without Ca2+.
- Argon is used to avoid oxidation.
- Glass slides are cleaned with Piranha solution.
Thank you all for your help, it's greatly appreciated!
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I first recommend you to read the following paper: Jass, J., Tjärnhage, T., & Puu, G. (2000). From liposomes to supported, planar bilayer structures on hydrophilic and hydrophobic surfaces: an atomic force microscopy study. Biophysical journal, 79(6), 3153-3163.
I noticed that you try to use DOPC. This is a typical bilayer forming lipid according to the structure-shape concept, see for example:
Cullis, P. T., & Kruijff, B. D. (1979). Lipid polymorphism and the functional roles of lipids in biological membranes. Biochimica et Biophysica Acta (BBA)-Reviews on Biomembranes, 559(4), 399-420. (free in Google Scholar, see figure 7)
Seddon, J. M. (1990). Structure of the inverted hexagonal (HII) phase, and non-lamellar phase transitions of lipids. Biochimica et Biophysica Acta (BBA)-Reviews on Biomembranes, 1031(1), 1-69. (see enclosed file)
My estimate would be that DOPC might be too stable, and you need a certain portion of so-called non-bilayer forming lipid.
Furthermore, in principle PC is considered as a neutral lipid, perhaps you need a certain portion of anionic (negatively charged) lipid.
My suggestion would be to first try a, in the literature, well-described mixture (in terms of composition and method) to make that work in your hands as well and then start to vary.
Hope this helps you somewhat.
Best regards.
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I collected daily feces from two mouse groups and stored them in -80°C. Now i'm prepared to compare TG content in feces of two groups following a method which was entitled "Lipid Extraction from Mouse Feces" (this method was published in Bio Protoc. 2015 Jan 5; 5(1): e1375.).
1. Dry the feces at 60°C overnight about 11 hours (this step is personally added)
2. Weigh exactly 1000mg of dry feces per mouse and powderize it using the tissue grinder.
3. Add 5ml of normal saline to 1000 mg of powderized feces in a 15-ml tube and vortex.
4. Add 5 ml of choroform: methanol mix (2:1 by volume)and vortex.
5. Centrifuge the suspension at 1000 × g for 10 min at room temperature. Afterwards a phenomenon that two liqiud phase seperated by a solid phase will happen. The lower liquid phase contains the extracted lipids in chlorofoem:methanol mix.
6.Use needle to drain out the lipid containg lower liquid phase.
7.Place the collected lower lipid phase in a stand under a fume hood, leave the tubes alone for 3-4 days until all liquid is evaporated.
8.Analyze lipid mass of feces in grams.
TO assess lipid concentration
9.Dissove solid lipid in 1ml isopropanol. Use Wako LabAssayTM Triglyceride(GPO ・ DAOS method) to test TG .
I have several questions and will
appreciate some advice from more experienced people.
Q1: After adding choroform: methanol mix (step4 ), how long will it takes to completely extract lipid from grined feces? Is it necessary to use a shaking platform for overnight?
Q2: when lower liqiuid phase drain out from the pore (step6), some granulum from solid phase may also come out which will greatly influence lipid mass analysis. Is there some advice?
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A period of overnight will be better for complete separation of both organic (Lipid) and inorganic layers. The flask should kept witout any disruption.
Filter the content (Organic-Lipid phase) using sodium sulphate to avoid the unwanted materials.
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For example the drug is highly hydrophillic and the lipid is hydrophobic in nature. When I add lysing solvents like ACN, methanol, tritonX100, the lipids are getting dissolved in the solvent. And since I need to estimate the drug content by LCMs method and the lipids should not be present in the sample solutions, I kindly request to advice the suitable method to do the separation.
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Try a water immiscible solvent to dissolve the lipids, add water if necessary, separate the aqueous layer and estimate.
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This dye itself has color of PE-Cy5 and after interaction with the lipid ROS, it emits green fluorescence. I have also found that this dye also interferes with the PE, PE-Cy7 channels somewhat. So I was wondering if there is an alternative to this dye? (Also, why is this the standard dye for ferroptosis related lipid peroxidation?)
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HNE
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So I am planning to make lipid nanoparticle with 3 different lipids. All I know is the final concentration of lipid is 2 mg/ml and the molar ratio (%) of A:B:C is 60:35:5.
Let's say the molecular weight for A = 550 g/mol, B = 700 g/mol, C = 2800 g/mol
How do I calculate the weight that I need for each components?
Thank you very much for the help
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Thank you very much Frederic and Noel, that helps a lot.
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the role of phytochemicals on cardiovascular diseases in specific mention to atherosclerosis, lipid profile
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In my experiment, I want to use phospholipid (Phosphatidic Acid) in my cell culture media. According to protocol, I soluted Phosphatidic Acid (PA) in chloroform then later diluted in culture medium and dispersed via 10 min of sonication in a water bath. But the solution is not clear and there are visible lipid layer and powder in surface. After vortex, it becomes turbid. If its property of phospholipid solutions? If not so, how can I get the clear solution for use in my experiment?
Looking for a kind response. Thank you
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Thanks Rob Keller Your guidelines helped me a lot to conduct my experiment smoothly
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Hi,
I am formulating liposomes (DSPC,DSPE-PEG200 and cholesterol) loaded with 5, 6 carboxyfluorescein (FAM). Briefly i rehydrated the dried lipid film with 5mM FAM dissolved first in 100 microL DMSO (to dissolve FAM fully) and added PBS (pH 7.4) up to a final volume of 1ml.
Since EE% is: (amount of FAM in sample/total amount added)*100. My question is how do i measure the exact encapsulation efficiency for FAM in my liposomes giving that there will be some FAM lost with purification step (i.e dialysis, column chromatography and extrusion) and also the volume collected after 3h (incubated at 55 degree with shaking at 160rpm) was less than the initial rehydration volume that was added into the lipid film.
Also will the lipids (PC, PE and Chol) interfere with the FAM readings in UV/vis spec (after disrupting them with an organic solvent)? or is it best to use Triton x100 to disrupt the vesicles?
Thanks and apologies for the long question.
Aziz
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Hi,
Abdulaziz Alobaid Abdulaziz Alobaid
Fluorescein amidites (FAM) encapsulated in lipid vesicles can be quantified using a protocol described in a nostalgic paper, attached.
Chen, R. F., & Knutson, J. R. (1988). Mechanism of fluorescence concentration quenching of carboxyfluorescein in liposomes: energy transfer to nonfluorescent dimers. Analytical biochemistry, 172(1), 61-77.
For more readings, have a look at these relevant papers:
Weinstein, J. N., Ralston, E., Leserman, L. D., Klausner, R. D., Dragsten, P., Henkart, P., & Blumenthal, R. (2018). Self-quenching of carboxyfluorescein fluorescence: uses in studying liposome stability and liposome-cell interaction. In Liposome technology (pp. 183-204). CRC Press.
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Ralston, E., Hjelmeland, L. M., Klausner, R. D., Weinstein, J. N., & Blumenthal, R. (1981). Carboxyfluorescein as a probe for liposome-cell interactions effect of impurities, and purification of the dye. Biochimica et Biophysica Acta (BBA)-Biomembranes, 649(1), 133-137.
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I would like to study the apo form (lipid-free) of a protein that only has been crystallized with lipids. I want to explore if it is possible to generate with a molecular dynamic a reasonable structure, making subtraction of lipids in several steps until obtaining the apo form. Likewise, I don't know if, during the molecular dynamic trajectory, it is possible to disappear lipids. I am thinking of using programs like GROMACS, AMBER, etc.
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You need to remove lipid before MD simulation. You can not delete or add any atom/residue/molecule during and after MD simulation, as it will destroy your trajectory data.
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What will be the ideal statistical test to compare fasting and random lipid levels in a nationwide cross-sectional study?
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Follow this article doi: 10.4103/1110-2098.215443
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Anyone with experience dissolving barite for intracrystalline lipids?
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There are traditional ways to extract lipids (keep in mind that lipids can mean several groups of compounds like fatty acids, cholesterol, phospholipids etc.). Roughly two are the most often used:
Bligh, E. G., & Dyer, W. J. (1959). A rapid method of total lipid extraction and purification. Canadian journal of biochemistry and physiology, 37(8), 911-917. https://www.tabaslab.com/protocols/BlighDyer.pdf
Folch, J., Lees, M., & Stanley, G. S. (1957). A simple method for the isolation and purification of total lipides from animal tissues. Journal of biological chemistry, 226(1), 497-509.
In your specific case you can find inspiration in:
Fatty acids in sparry calcite fracture fills and microsparite cement of septarian diagenetic concretions
Blyth, A. J., Farrimond, P., & Jones, M. (2006). An optimised method for the extraction and analysis of lipid biomarkers from stalagmites. Organic Geochemistry, 37(8), 882-890. https://doi.org/10.1016/j.orggeochem.2006.05.003(see enclosed file)
Hope this helps you a bit further.
Best regards.
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I am preparing liposomes using the thin film hydration method. The lipid I am using is DPhPC (no phase transition temperature between -120/ 120 C). Briefly here is my protocol
I first dissolve the lipid (only DPhPC) in Chloroform (1 mg/ml) using a round flask. Then I dry the chloroform with nitrogen. The dried lipid film is then placed in a vacuum chamber for 8+ hrs. Afterwards, 1x PBS solution (1 ml) is added to the flask and the solution is allowed to hydrate at a temperature of 65 ⸰ C under vigorous stirring with a magnetic spin bar for 2+ hrs.
My problem, is that after chloroform evaporation, a non uniform white residue appears on the wall instead of a thin uniform lipid layer. Also, during hydration the dried lipid is not breaking into smaller pieces and remain as large aggregates (see image attached) which is not possible to extrude and sonication does not help. Any idea on why hydration is not properly occurring?
note: The attached image is taken after 2 hrs of hydration, it shows the large precipitate of lipids in 1xpbs.
Thank you in advance,
Rami
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First of all, how fresh is your DPhPc?
Properties of diphytanoyl phospholipids at the air-water interface by Yasmann and Sukharev (DOI: 10.1021/la503800g) use a similar method to prepare DPhPc liposomes with some slight, but significant differences. Consider them when comparing with your methods.
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I want to extract lipid and protein from mice whole serum followed by quantification of Sialic acid in total fat. I found many methods but I am wondering, is there any kit available?
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Because it is a nonionic detergent i do not think it will effect the protein.
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I have an MDS trajectory for a lipid bilayer comprising of POPC, POPE, POPSm cholesterol, and sphingomyelin molecules. the system was built using CHARMM-GUI and equilibrated in NAMD and area per lipid was calculated. How can I know that this value indicates that the system was equilibrated? what are the reference values for area per lipid?
Thanks
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Hello,
I think we usually consider an average of 60-70Ų. (I always use 70 as a proxy when I calculate the number of phospholipids per layer, per liposomes of a give size). Depending on the lipid of interest and the membrane properties, this can vary between 40 and 70 (according to the following paper)
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In preparing a liposome loaded with an aqueous extract to obtain a powdered sample, is it recommended to dissolve the extract in distilled water, then use same to hydrate the already formed lipid film? Thank you.
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Thank you Mohanad.
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I am working on the disruption of some yeast genes to create new strains... for my research project "Metabolic Engineering of Saccharomyces cerevisiae as lipid cell factories for Bio-Diesel production"
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While you can use point mutations to introduce premature stop codons close to the start of the open reading frame, this would leave room for revertants to occur. As such, deletion of the genes in question is safer. See
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I am currently extracting crude lipids in musa(banana) via soxhlet extraction using petroleum ether. I wanted to know if anyone had better yield via the acid hydrolysis method.
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For crude lipid/fat, Sohxlet would be better choice. Whereas, use a combination of organic solvents for the estimation of total lipids.
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Hello!
How can the total amount of lipids in the liver be biochemically determined without the use of chloroform and methanol?
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The relationship between lipid profile and severity of liver damage in cirrhotic patients
This article might help you.
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Dear colleagues,
I have a study data from a sample population that contains three different subsets of study subjects: 1. Healthy control with normal BMI, 2. Non-diabetic overweight/obese subjects and 3. Type 2 diabetic subjects.
I am trying to explore whether plasma lipid parameters, their ratios or indices could potentially predict the incident metabolic syndrome in the given sample population (N=142). For this, I want to develop a linear regression model using metabolic syndrome components (expressed in number as 0, 1, 2, 3, 4,and 5) as the dependent variable and all plasma lipids, their ratios and indices as the independent variables. My question is how should I develop this regression model? Should I develop the model using the data of whole study population that contains all three subsets of subjects mentioned above or first split the whole study data into two groups as " with metabolic syndrome" and " without metabolic syndrome" apply linear regression and report the beta coefficient values of only the subset of " with metabolic syndrome"?
Please also advise me whether there could another regression method that could best apply to my case. Thank you.
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Thank you Tuyen Van Duong for clearing my confusion.
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Considering DNA/RNA viruses, naked RNA (viroids) and proteins (prions) which are believed to be causative agents of particular severe infectious diseases, is there any (theoretical) infectious potential for the rest of biological macromolecules (lipids and polysacharides)? Just interested if someone explored this possibility and if there are some papers dealing with this idea (I was unable to find any). Thanks for your suggestions :)
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Hi Martin. As Vaclav says, polysaccharides and lipids can be immunostimulatory/modulatory. For example, bacterial lipopolysaccharide and glucans in the cell wall of fungal pathogens are key pathogen-associated molecular patterns. My background is in the host recognition of polysaccharides in pathogens, so I'm happy to discuss if I can provide any guidance.
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Hi friends,
I am looking to find out the top scientists and their clusters who are working in the field of Hyperlipidemia, Obesity and Lipid research. Please suggest me best methods to find out.
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I agree with Kabelo Mokgalaboni.
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Hi
I want to do a simulation of a lipid bilayer representing the lipid membrane of cancer cells.
What are the proportions of different lipids that I should use in my system?
Thanks
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In the matini official website, On matini's website, many lipids don't have mapping files. And i want to edit a mapping file for a lipid molecular by myself, could you help me to solve the problem?
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So beta-oxidation is a proces in which fatty acids are broken down and results in two molecules, where one is Acetyl-CoA. However, if I look at the the scheme of beta-oxidation, I don't understand which is/are the fatty acids in this process. So for example, Acyl-CoA will eventually result in myristoyl-CoA and Acetyl-CoA. What is the fatty acid in this beta-oxidation process?
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The long-chain acyl-CoA enters the fatty acid β-oxidation pathway resulting s in the production of one acetyl-CoA from each cycle of fatty acid β-oxidation.
β-Oxidation is the major metabolic pathway by which energy is released from fatty acids.
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Hi all,
I have tried to do a calcein leakage assay by LUVs that I made (dopc:dopg 1:3, dopc:dopg 3:1, and dopc only). In doing so I dissolved my lipids film in 2 mL of 10mM tris buffer containing 70mM calcein and after incubation at RT for 1h and freez-thaw for 5 cycles, I extrude them by a mini extruder with 10mm filter support and 0.1um PC membrane. However, after centrifugation I got no pellet. But when I skipped the extrusion step and directly centrifuged my samples I got pellet. Thus, I believe my lipids are missed during the extrusion but I have no idea how should I overcome this issue. Does anyone have any idea?
Cheers,
Maryam
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Hi Maryam,
It happens that a portion of your lipids gets stuck in your extruder. It also happens that some volume of the whole solution stays in the extruder, especially when the extruder is dry before usage.
Before extrusion, I always extrude an empty buffer a few times to fill in all the small voids in the extruder with the buffer and not to lose the sample. With this step, I also check if everything is tight and I do not have any spill.
Also, make sure that your extruder is properly cleaned before you use it. There might be some leftover molecules from the previous usage, which stick to your lipids and make it impossible to go through. TO properly clean it, use methanol multiple times, then use chloroform for the glass and metal parts, and then put everything to the sonication bath. You can sonicate it first in methanol, second in ethanol, and third in water right before you use it again to get rid of any organic solvents left in there. If there are organic solvents in the extruder, they will dissolve your lipids.
That's everything that comes into my mind. Good luck in making it work!
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Hi!
I am experiencing some difficulties in preparing brain tissue for proteomic analysis - most likely due to high lipid content that interferes with proteomic data acquisition.
We need to isolate proteins in their native state including larger molecular weight complexes and protein aggregates. This means that we are limited to bead beating in native lysis buffer (1 mM MgCl2, 150 mM KCl, 100 mM HEPES, pH 7.4) and a very mild centrifugation step (800xg, 5 min) before using the supernatant for tryptic digest, C18-cleanup and HPLC-MS.
However, I commonly observe column clogging or ion suppression using these samples, which I suspect is due to the presence of contaminating lipids.
Does anyone have a good protocol to remove lipids without denaturing proteins or for removing lipids after the tryptic digest?
Any advice is highly appreciated! Many thanks in advance!
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Dear Norbert,
Your question is a challenging one!...and as you say the clogging and ion suppression issues in the LC-MS analysis may be due to the presence of lipids in your sample.
As removal (extraction) of lipids typically involves the use of organic solvents this will also denature your protein. So in my opinion, it might be an option to remove lipids after the tryptic digestion step (and assuming that lipids do not restrict the protein digestion).
You can find commercially available SPE columns (Supelco) designed to retain (phospho)lipids from biological samples - SPE Hybrid-Phospholipid.
Hope this helps,
Ana
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Hi,
One of my protein of interest exhibits a lipid peroxidase activity. I have solved the X-ray structure of the protein alone which has a hydrophobic channel.
I’m looking for a software adapted to docking driven by hydrophobic interactions.
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Autodock software is good for docking lipids to proteins
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I currently store my 10 mg/mL mixed composition of POPC+PGPC (9:1 mol/mol) in a chloroform/methanol (9:1, v/v) solution. For the BLM experiment I require a working volume of ~40-50 uL of this solution which I dry into a lipid film and subsequently re-suspend in organic solvents. I use a decane : butyl alcohol mix for plain POPC, but I am having trouble with stabilizing my POPC+PGPC due to PGPC's highly oxidized nature.
So far I have tried (v/v) 50:1, 20:1, 10:1, 5:1, and 3:1 ratios of decane to butyl alcohol, and in each case my lipid precipitated into a milky substance from between 30 minutes to 1.5 hours.
I would greatly appreciate it if someone had any insight into how I could stabilize my lipids in organic solvent for an entire day? No papers on supported lipid bilayers/BLM have insight into working with oxidized lipids.
Thank you, in advance.
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@Yuri Mirgorod @Michal Surma
Hello, again;
I thought I would update you. It turns out that the POPC we had ordered was faulty (old, or manufacturing error). We purchased new POPC and my lipids stayed in solution.
It is worth noting that I have had much better results when ordering lipids that are pre-dissolved in chloroform, rather than opening the ampule and adding chloroform once they arrive in the lab.
Thank you for your input, again.
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The metabolic syndrome is characterized by dyslipidemia and hypertension in obese individuals. Lipoprotein subfractionation of plasma from obese individuals indicate the presence of large very low density lipoprotein, small dense low density lipoprotein and decreased high density lipoprotein. Other defects in obesity and Alzheimer’s disease may be linked to specific lipid species. In obesity and Alzheimer’s disease toxic ceramides are found in these chronic diseases and linked to inflammation, insulin resistance and amyloidogenesis. The search for lipids associated with longevity has accelerated to extend lifespan in the developing and developed world and specific lipids by LIPIDOMICS in the plasma need to be identified to determine the lipids involved in cell signalling associated with healthy aging.
RELEVANT REFERENCES:
1. Victor Bustos and Linda Partridge. Good ol’ fat: links between lipid signaling and longevity. Trends Biochem Sci. 2017 Oct; 42(10): 812–823.
2. Duffy J., Mutlu A.S., Wang M.C. (2017) Lipid Metabolism, Lipid Signalling and Longevity. In: Olsen A., Gill M. (eds) Ageing: Lessons from C. elegans. Healthy Ageing and Longevity. Springer, Cham.
3. Constantinou, J.K., Southam, A.D., Kvist, J. et al. Characterisation of the dynamic nature of lipids throughout the lifespan of genetically identical female and male Daphnia magna. Sci Rep 10, 5576 (2020).
4. Pradas et al. Exceptional human longevity is associated with a specific plasma phenotype of ether lipids. Redox Biology. Volume 21, February 2019, 101127
5. Lim WLF, Huynh K, Chatterjee P, et al. Relationships Between Plasma Lipids Species, Gender, Risk Factors, and Alzheimer's Disease [published online ahead of print, 2020 May 26]. J Alzheimers Dis. 2020;10.3233/JAD-191304. doi:10.3233/JAD-191304.
6. W. L.Florence Lim, I.J. Martins, R.N.Martins. The involvement of lipids in Alzheimer's disease. J Genet Genomics. 2014; 41(5):261-74.
7. Martins IJ, Creegan R, Lim WLF and Martins RN. (2013). Molecular insights into appetite control and neuroendocrine disease as risk factors for chronic diseases in Western countries. Special Issue. Molecular Mechanisms Involved in Inflammation and Insulin Resistance in Chronic Diseases and Possible Interventions. Open Journal of Endocrine and Metabolic Diseases. 3;11-33.
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Here are some papers:
Our comparative genetic study in humans and C. elegans suggests lipid/lipoprotein metab is a major risk for AD:
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I am planning to prepare liposomes with DMPC and phytosterols using the thin film method to encapsulate a hydrophobic drug. Usually, after the evaporation of the lipids in a vacuum assisted rotary evaporator, an additional drying step is perform to eliminate traces of solvent from the lipid film. Does flushing with an inert gas (such as Nitrogen) holds an advantage over drying in a vacuum desiccator?
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Dear Swee Liang Kho it depends on how much sample you have. For larger amount of lipid, the last traces of solvent can be best removed by placing the sample under a high vacuum overnight. Smaller samples of lipid solution (less than ca. 1 mL) can easily be dried under a stream of nitrogen. Thus in the end both methods are useful. In this context, please have a look at the following useful ink:
Making Liposomes in the Lab
Yet another potentially helpful link is the following:
How do I remove solvent from a small amount of lipid?