FAQ

Commonly used solvents include methanol, acetonitrile, and water, either alone or in various combinations. Methanol is particularly effective for extracting a broad range of metabolites due to its ability to solubilize both polar and non-polar compounds. A common extraction protocol involves using a mixture of methanol and water to precipitate proteins while solubilizing metabolites. Acetonitrile is another excellent choice, especially for polar metabolites, as it can efficiently extract compounds without significantly disrupting the sample matrix. Some methods may even employ a two-phase extraction, where the sample is treated with both aqueous and organic solvents to separate different classes of metabolites. The choice of solvent should align with the specific metabolites of interest and the intended analytical methods to ensure optimal extraction efficiency and reproducibility.

To ensure complete extraction of metabolites from tissue samples, it's important to optimize several factors, including the extraction method, solvent choice, and processing conditions. For instance, using a suitable solvent that matches the polarity of the target metabolites is key. Additionally, methods like homogenization can help break down tissue structures, making metabolites more accessible to solvents. Techniques such as sonication or mechanical grinding can significantly enhance extraction efficiency. Furthermore, controlling extraction time and temperature is crucial. Longer extraction times can lead to better recovery, but they may also cause degradation of unstable metabolites.

Yes, spiking metabolite standards into samples before extraction is a common practice for quantification purposes. This approach allows us to create calibration curves that can be used to quantify the concentration of metabolites in unknown samples. By adding known amounts of standards to the samples, we can account for variations in recovery and extraction efficiency, ultimately improving the accuracy of their quantification. However, it’s important to consider the timing and conditions under which the standards are added. For instance, adding standards too late in the extraction process may not accurately reflect their behavior during extraction. Ideally, standards should be spiked into the sample prior to any processing steps to mimic the actual extraction conditions and ensure that their recovery aligns with that of the metabolites of interest.

Protein precipitation is a widely used technique in metabolomics to prepare biological samples for analysis. This method involves adding an organic solvent, such as methanol or acetonitrile, to the sample to precipitate proteins. The rationale is that proteins can interfere with downstream analyses, so removing them can help enhance the clarity of the metabolomic profile. After adding the solvent, the mixture is typically centrifuged to separate the precipitated proteins from the supernatant, which contains the metabolites. This step is crucial, as it allows researchers to concentrate the metabolite fraction while discarding potential contaminants.

Solvent temperature can significantly influence the efficiency of metabolite extraction. Generally, higher temperatures can increase the solubility of metabolites, leading to improved extraction yields. Warm solvents may facilitate better diffusion of metabolites from the sample matrix into the solvent, enhancing recovery rates. However, this must be balanced against the stability of the metabolites, as some compounds may degrade or alter at elevated temperatures. Conversely, using cold solvents can be beneficial for preserving sensitive metabolites that are prone to degradation or reaction at higher temperatures. In these cases, lower temperatures can help stabilize the metabolites during the extraction process.

While solid-phase extraction (SPE) is a powerful tool for metabolomics, there are several common pitfalls that researchers should be aware of. One major issue is the potential for incomplete elution of target metabolites, which can lead to underestimations of their concentrations. This can occur if the chosen sorbent is not well-suited for the specific metabolites or if the elution conditions (e.g., solvent choice or volume) are not optimized. Another challenge is the introduction of bias during sample handling. For example, if samples are not properly equilibrated with the sorbent or if the flow rate is too fast, it can affect retention and recovery. Contaminants from the sorbent itself can also interfere with the analysis, making method validation and careful selection of SPE cartridges essential for reliable results.

Yes, different extraction methods can indeed lead to biases in the metabolic profile obtained from samples. Each extraction technique is designed to target specific classes of metabolites, and if the method does not adequately capture all relevant metabolites, the resulting profile may be skewed. For example, using a non-polar solvent for extraction may result in the loss of polar metabolites, leading to an incomplete representation of the metabolic state. Additionally, different extraction conditions, such as temperature, time, and agitation, can affect the recovery rates of certain metabolites differently. These variances can introduce biases, making it difficult to compare metabolic profiles across studies that employ different methods. Therefore, we must carefully choose and standardize extraction techniques to ensure consistency and reproducibility in their analyses.

Extracting volatile organic compounds (VOCs) requires specialized methods to capture their transient nature. One common technique is headspace sampling, where the air above a sample is analyzed for VOCs. In this method, samples are sealed in a vial, and the air above the liquid or solid phase is allowed to equilibrate, allowing VOCs to partition into the gas phase. This gas can then be collected and analyzed, often using techniques like gas chromatography-mass spectrometry (GC-MS). Another method involves using solid-phase microextraction (SPME), where a fiber coated with a sorbent is exposed to the sample to adsorb VOCs. After a designated time, the fiber is then desorbed into a GC-MS for analysis. These methods are particularly effective for capturing and analyzing VOCs, which are often present in low concentrations but can provide valuable insights into metabolic processes and environmental interactions.

One common method is using organic solvents, such as methanol or ethanol, to disrupt the cell membrane and solubilize intracellular metabolites. This is often followed by centrifugation to separate the cellular debris from the supernatant, which contains the metabolites. The methanol-water extraction is frequently employed because it effectively captures a wide range of polar and semi-polar metabolites. Another approach is solid-phase extraction (SPE), which can further purify the extracts after initial solvent extraction. This method can help concentrate metabolites and remove impurities, making it particularly useful for downstream analysis. Additionally, some researchers use ultrasound-assisted extraction to enhance the efficiency of metabolite recovery from microbial cells. By applying ultrasonic waves, this technique can facilitate better penetration of the solvent into the microbial matrix, leading to higher yields.

The choice of extraction buffer can significantly impact downstream metabolomic analysis by influencing metabolite stability, solubility, and recovery. Different buffers have varying pH levels and ionic strengths, which can affect the extraction efficiency of specific metabolites. For instance, a phosphate buffer at physiological pH might be ideal for extracting amino acids, while a more acidic buffer might be necessary for certain organic acids. Moreover, the presence of salts or additives in the buffer can either enhance or inhibit the detection of metabolites in mass spectrometry. Some compounds may precipitate or degrade in certain buffers, leading to biases in the metabolomic profile.

Yes, simultaneous extraction of metabolites and proteins can be achieved, although it requires careful optimization of extraction conditions. One common approach is the use of protein precipitation methods combined with organic solvent extraction. For instance, adding methanol or acetonitrile to a biological sample can precipitate proteins while simultaneously solubilizing metabolites, allowing both to be extracted in a single step. However, this method must be optimized to ensure that the recovery of both metabolites and proteins is adequate. Factors like solvent volume, temperature, and extraction time can influence the efficiency of the extraction.

Extraction efficiency can vary significantly with different solvents due to their polarity and ability to solubilize specific classes of metabolites. For example, methanol and ethanol are polar solvents that are effective at extracting a wide range of hydrophilic metabolites, including amino acids and organic acids. They tend to work well for cellular components due to their ability to penetrate biological membranes. On the other hand, chloroform is a non-polar solvent commonly used in liquid-liquid extraction methods to capture lipids and hydrophobic metabolites. Water, while excellent for very polar compounds, may not extract non-polar metabolites effectively.

Methods for extraction of hydrophilic compounds: Sample was thawed on ice. Take 50 ± 2 mg of one sample and add cold steel balls to the mixture and homogenate at 30 Hz for 3 min. Add 1 mL 70% methanol with internal standard extract to the homogenized centrifuge tube, Whirl the mixture for 5 min and stand on ice for 15 min, then centrifuge it with 12,000 rpm at 4 ℃ for 10 min. After centrifugation, draw 400 uL supernatant into the corresponding EP tube and store in -20 ℃ refrigerator overnight, then centrifuge at 12,000 rpm at 4 ℃ for 3 min, and take 200 uL supernatant to the liner of the corresponding injection bottle for on-board analysis.

We can concentrate it by lyophilizer.

For Cell sample 1) Methods for extraction of hydrophilic compounds Sample was thawed on ice, then added 1 mL pre-cooled extractant (80% methanol aqueous solution), and whirl for 2 min. Freeze the mixture for 5 min in liquid nitrogen after remove ice for 5 min, it will be whirled for 2 min, circulate this at 3 times. Centrifuge the mixture again with 12,000 rpm at 4 ℃ for 10 min. Finally take 200 uL supernatant to the liner of the corresponding injection bottle for on-board analysis. 2) Methods for extraction of hydrophobic compounds Sample placed in liquid nitrogen for 2 min，then thawed on ice for 5 min and vortexed homogenously. Repeat the first step 3 times, then centrifuge it with 5,000 rpm at 4 ℃ for 1 min. Homogenize it with 1mL extractant (include methanol，MTBE and internal standard mixture). Vortex the mixture for 2 min, sonicate for 5 min. Then add 200 uL water and whirl the mixture for 1 min, and centrifuge it with 12,000 rpm at 4 ℃ for 10 min. Extract 500 uL supernatant and concentrate it. Dissolve powder with 200 uL mobile phase B, then stored in -80 ℃. Finally take the dissolving solution into the sample bottle for LC-MS/MS analysis.

Plasma metabolites extraction for untargeted metabolomics: The sample stored at -80 °C refrigerator was thawed on ice and vortexed for 10 s. 50 μL of sample and 300 μL of extraction solution (ACN : Methanol = 1:4, V/V) containing internal standards were added into a 2 mL microcentrifugetube. The sample was vortexed for 3 min and then centrifuged at 12000 rpm for 10 min (4 °C). 200 μL of the supernatant was collected and placed in -20 °C for 30 min, and then centrifuged at 12000 rpm for 3 min (4 °C). A 180 μL aliquots of supernatant were transferred for LC-MS analysis.

We possess extensive experience in processing tear samples collected using Schirmer strips, and we will provide detailed information on this process later. For tear samples, we have project experiences covering untargeted metabolomics, TM widely-targeted metabolomics and quantitative lipidomics. Besides, we also have the experience in neurotransmitter-targeted metabolomics for tears.

The samples were extacted by 100% methanol as provided. we can do Widely-Targeted Metabolomics for Plants.Normally, 50mg samples with 1 ml methanol were used for extraction. After concentration, 100ul will be fine

The extraction reagent used for energy metabolism detection contains certain organic agents, which can cause protein denaturation and precipitation. We will use liquid nitrogen for repeated freeze-thaw cycles on the samples to rupture the cells, allowing the metabolites to be fully dissolved. We use the external standard method for quantification of energy targeted metabolomics detection. Internal standards only participate in quality control, used to evaluate whether the preprocessing and instrument operation are abnormal after the samples arrive at the laboratory. Therefore, energy metabolism assay does not detect internal standard recovery levels. When developing methods, we compare the effects of different extraction reagents and their proportions on extraction efficiency. Currently, the extraction method we use, which consists of 500 μL 80% methanol, is the optimal condition after comparison. Some literature mentions extraction methods involving methanol/acetonitrile/water (2:2:1) and methanol/acetonitrile/water (5:3:2). We have also compared them, and there is no significant difference in extraction efficiency compared to 80% methanol.

It is challenging to remove the metabolites interference from the FBS contained in the media. We should provide an adequate number of blank controls so that we can calculate the actual content of metabolites secreted from cells into media through differential analysis.

We can process the cell-free supernatant (CFS) extracts and bacterial cells mentioned by the client using our metabolomics assays. For the CFS extracts, a minimum of 20 mg of sample is required, but we recommend providing 100 mg for optimal results. For bacterial cells, lyophilization is not necessary. Simply collect the cells, wash them with PBS, and store the cell pellets at -80°C. We recommend a cell count of 10^10 cells per sample. For the MRS medium control, we suggest providing 100 µl of sample.