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0.1 M sodium citrate in 10% ethanol and DNA solubility

0.1 M sodium citrate in 10% ethanol and DNA solubility



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I understand how DNA precipitation works in the presence of salt (such as 0.3 M sodium acetate or 0.2 M sodium chloride) and alcohol (30 ~ 50 % isopropanol or 60 ~ 75% ethanol)

However, in the Trizol kit manual, DNA could be washed and precipitated in0.1 M sodium citrate in 10% ethanol

At only 10% ethanol and 0.1 M sodium ion, how can DNA be in a "compact/insoluble" form so it can be precipitated by a low speed centrifugation (2000 x g for 5 mins)?


The sodium ions will bind to and neutralize the (-) charge on the phospho backbone of DNA. This reduces hydrogen bonding sites for water, which then results in less solubility.

Edited for clarity and because I didn't answer the question the first time. Sorry about that.

The extraction of DNA is from the interphase layer, which is a very small fraction of the original sample. The extraction of the interphase layer is then mixed with 100% EtOH (300 ul per 1.0 Trizol used in that sample). This should actually be 60% or more EtOH based on my experience using this method.

That is the precipitation step.

The fact that the wash has only 10% EtOH and 0.1M sodium citrate and that is still sufficient to keep DNA precipitated revolves around 0.1M sodium citrate is usually 0.3M sodium ions (sodium citrate can be mono-, di- or tri-sodium, but unless specified… sodium citrate implies tri-). Remember for the wash, you don't need to do a quick precipitation, you just need to prevent it from solubilizing again.

I am removing my comments that contained the same information.


The single-step method of RNA isolation by acid guanidinium thiocyanate–phenol–chloroform extraction: twenty-something years on

Since its introduction, the 'single-step' method has become widely used for isolating total RNA from biological samples of different sources. The principle at the basis of the method is that RNA is separated from DNA after extraction with an acidic solution containing guanidinium thiocyanate, sodium acetate, phenol and chloroform, followed by centrifugation. Under acidic conditions, total RNA remains in the upper aqueous phase, while most of DNA and proteins remain either in the interphase or in the lower organic phase. Total RNA is then recovered by precipitation with isopropanol and can be used for several applications. The original protocol, enabling the isolation of RNA from cells and tissues in less than 4 hours, greatly advanced the analysis of gene expression in plant and animal models as well as in pathological samples, as demonstrated by the overwhelming number of citations the paper gained over 20 years.


Protocol - TRI Reagent DNA Isolation

DNA is precipitated from the phenol phase and interphase of samples that have been homogenized (or lysed) in 1 ml of TRI Reagent (step 5 in the RNA Isolation Protocol). After a series of washes to remove residual phenol, the DNA pellet is solubilized in a mild alkaline solution, and the pH is adjusted. This technique performs well with samples containing >10 μg of DNA.

  • 100% ethanol, ACS grade or better
  • Nuclease-free water
  • Trisodium citrate
  • NaOH
  • HEPES (free acid)
  • Appropriately sized nuclease-free centrifuge tubes with secure closures, compatible with phenol/chloroform, and capable of withstanding centrifugal forces of 12,000 x g.
  • DNA Wash Solution: 0.1 M trisodium citrate in 10% ethanol (no pH adjustment required), 2–3 ml per 1 ml of TRI Reagent used in the initial homogenization:
  • 75% ethanol, 1.5–2 ml per 1 ml TRI Reagent used in the initial homogenization
  • 8 mM NaOH,300–600 μl per 50–70 mg tissue or 10 7 cells
  • 0.1 M or 1 M HEPES (free acid), see Table 1
  • Unless stated otherwise, conduct the procedure at room temperature.
  • The molecular weight of the recovered DNA depends on the shearing forces applied during homogenization. If recovery of high molecular weight DNA is desired, use a loosely fitting homogenizer in the initial homogenization step of the RNA Isolation Protocol. Avoid using a Polytron homogenizer.
  • If the DNA is isolated only for quantitative purposes: a) samples can be more vigorously homogenized, including the use of a Polytron b) the phenol phase and interphase can be stored at 4°C for a few days or at –70°C for a few months c) the DNA can be solubilized using 40 mM NaOH instead of an 8 mM solution, and by vortexing the DNA pellet instead of pipetting.

The starting material for this procedure contains TRI Reagent, which contains a poison (phenol) and an irritant (guanidine thiocyanate). Contact with TRI Reagent will cause burns and can be fatal. Use gloves and other personal protection when working with TRI Reagent.


Neurobiology of Cytokines

Dan Lindholm , . Eero Ċastrén , in Methods in Neurosciences , 1993

Solutions

Buffer D (denaturing buffer):

Guanidium thiocyanate (4 M)

Sodium citrate (25 mM), pH 7

Sarkosyl lauryl sulfate (0.5%)

Mix guanidium, sodium citrate, and sarkosyl, and store in 50-ml Falcon tubes.

Before use, add 360 μl of 2-mercaptoethanol/50 ml.

Chloroform-isoamyl alcohol, 49:1

ComponentStockFor 300 μlFor 600 μl
NaHPO4 (100 mM), pH 7100 mM30 μl60 μl
Deionized glyoxal 70 μl140 μl
Dimethyl sulfoxide 200 μl400 μl
ComponentStockFor 20 mlFor 40 ml
Deionized formamide (50%)100%10 ml20 ml
NaHPO4 buffer (50 mM), pH 71 M1 ml2 ml
SSC (3×)20×6 ml12 ml
SDS (0.5%)10%1 ml2 ml
Na2EDTA (5 mM)0.5 M200 μl400 μl
ssDNA (250 μg/ml)10 mg/ml500 μl1 ml
Denhardt's solution (5×)50×2 ml4 ml

Enzymatic bioconversion of citrus hesperidin by Aspergillus sojae naringinase: Enhanced solubility of hesperetin-7-O-glucoside with in vitro inhibition of human intestinal maltase, HMG-CoA reductase, and growth of Helicobacter pylori

Hesperetin-7-O-glucoside (Hes-7-G) was produced by the enzymatic conversion of hesperidin by Aspergillus sojae naringinase due to the removal of the terminal rhamnose. Extracts from orange juice and peel containing the hesperidin were so treated by this enzyme that the hesperidin could also be converted to Hes-7-G. The solubility of Hes-7-G in 10% ethanol was enhanced 55- and 88-fold over those of hesperidin and hesperetin, respectively, which may make Hes-7-G more bioavailable. Hes-7-G was 1.7- and 2.4-fold better than hesperidin and hesperetin, respectively, in the inhibition of human intestinal maltase. Hes-7-G was more potent by 2- and 4-fold than hesperidin in the inhibition of human HMG-CoA reductase. Additionally, Hes-7-G exhibited more effective inhibition of the growth of Helicobacter pylori than hesperetin, while its effectiveness was similar to that of hesperidin. Therefore, the results suggest that bioconverted Hes-7-G is more effective and bioavailable than hesperidin, as it has enhanced inhibitory and solubility properties.

Highlights

► Hes-7-G could be converted from hesperidin by Aspergillus sojae naringinase. ► Hes-7-G was highly more soluble than hesperidin and hesperetin. ► Hes-7-G was more potent than hesperidin in the inhibition of intestinal maltase. ► Hes-7-G was more potent than hesperidin in the inhibition of HMG-CoA reductase.


Methods

Ethics Statement

This study was carried out in accordance with the guidelines approved by the Committee on Human Care and Use by the NASA and JAXA Ethical Review Board and the Human Research Multilateral Review Board (HRMRB). All participants provided written informed consent.

Hair sample preparation

Ten astronauts at the International Space Station (ISS) participated in the study. They were at the ISS on a 6-month-long mission. During each mission, five strands of hair were sampled six times from each astronaut. The 6 different sampling times were as follows (Fig. 1): first Preflight (Launch (L)-180

3 months before launch), second Preflight (L-60

14: from 2 months to 2 weeks before launch), first Inflight (L + 20

37: from 20 to 37 days after launch), second Inflight (Return (R)-20

7: from 20 117 to 7 days before return), first Postflight (R + 2

7: from 2 to 7 days after return), and second Postflight (R + 30

90: from 1 to 3 months after return). The sampling days differed for each astronaut because of differing schedules. In each mission, two astronauts were paired, and individual hair samples were collected. For each individual sample, five strands of hair were grasped as close as possible to the scalp and pulled out using tweezers in the direction of hair growth without damaging the hair roots. The Preflight and Postflight samples were stored at −80 °C until analysis. The Inflight samples were stored in the Minus Eighty Degree Laboratory Freezer on the ISS (MELFI) as soon as possible after collection until being returned for analysis.

RNA extraction

Hair roots (approximately 2–3 mm) were used as the source for extracted mRNA. The roots were cut into approximately 15 fragments (0.1–0.2 mm each) by using a microsurgical knife under a stereoscopic microscope. The collected fragments were immersed in 800 μl of ISOGEN reagent (Nippon Gene Toyama, Japan) in tubes and stirred (15 sec × 2 times) using a Bioruptor UCD-250 sonication device (Cosmo Bio Tokyo, Japan). Next, the RNA was purified from hair lysates using an ISOGEN kit according to the manufacturer’s instructions. Briefly, the tubes were maintained at room temperature for 5 min, followed by the addition of 200 μl of chloroform. The subsequent process of RNA purification was performed according to the manufacturer’s instructions. After isolation, RNA pellets were washed with 70% ethanol, air dried, and resuspended in 10 μl of RNA-free water (Gibco-BRL Gaithersburg, MD). Total RNA was quantified at 260 nm using a NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies Inc. Wilmington, DE). RNA quality was determined using an Agilent Bioanalyzer 2100 (Agilent Technologies Palo Alto, CA). The 28S:18S rRNA ratio and the RNA integrity number (RIN) were calculated using the 2100 Expert and RIN Beta Version software (Agilent Technologies), respectively.

RNA amplification

As the RNA sample extracted from the hair samples was small, a double RNA amplification step was incorporated prior to microarray hybridization. Total RNA was amplified using an Ambion MessageAmp aRNA Kit (Thermo Fisher Scientific Waltham MA, USA). Briefly, first- and second-strand cDNA were synthesized. Unlabeled aRNA was generated by in vitro transcription with non-biotinylated NTPs. For probe preparation, RNA was reverse-transcribed with second-round primers. The second-strand cDNA was synthesized with T7 oligo(dT) primers and purified. Biotin-labeled cRNA was generated via in vitro transcription and purified using an RNeasy Kit (Qiagen Venlo, Netherlands).

DNA extraction from hair RNA samples

DNA samples from the astronauts’ hair roots were extracted using ISOGEN (Nippon Gene Toyama, Japan) from the rest of the extracted RNA samples according to the manufacturer’s instructions. Briefly, 0.3 ml of 100% ethanol were added to each sample, followed by a 3-minute incubation at room temperature. Following centrifugation using a refrigerated microcentrifuge (Kubota model 3500 KUBOTA Tokyo, Japan) at 2000 g and 4 °C for 5 minutes, the precipitate in the tubes was washed with 1 ml of 0.1 M sodium citrate in 10% ethanol for 30 minutes. After further centrifugation at 2000 g and 4 °C for 5 minutes, precipitates were washed again with 1 ml of 0.1 M sodium citrate in 10% ethanol for another 30 minutes. The isolated DNA pellets were washed with 1 ml of ethanol and then centrifuged at 2000 g and 4 °C for 5 minutes. The rinsed DNA samples were air dried and re-suspended in 50 μl of Tris-ethylenediaminetetraacetic acid (EDTA) buffer solution (TE solution). To quantify the amount of DNA, the absorbance rates at 260 nm were measured using a U-1900 spectrophotometer (Hitachi High-Tech Science Tokyo, Japan). The 260 nm/280 nm absorbance ratio of the DNA samples was also measured to determine DNA quality.

Measurement of mtDNA/nDNA, mtRNA/nRNA and mtRNA/mtDNA ratios

The ratio of mitochondrial (mt) and nuclear (n) DNA and the ratio of mtRNA and nRNA from the astronauts’ hair roots were analyzed via SYBR Green-based quantitative PCR (qPCR). The DNA and RNA samples described above were used for qPCR. Ten nanograms of DNA were used to detect mt or nDNA, and 2 ng of RNA was used to detect mt or nRNA. The qPCR reactions were performed with the ABI Prism 7000 sequence detection system (Applied Biosystems Foster City, CA, USA) using a QuantiTect SYBR Green PCR Kit (QIAGEN Valencia, CA, USA) as recommended by the manufacturer, except for the following modifications. For ND1 and GAPDH amplification, the denature temperature was 95 °C, the denature time was 15 seconds, the annealing temperature was 55 °C, the annealing time was 30 seconds, the elongation temperature was 72 °C and the elongation time was 40 seconds.

NADH dehydrogenase subunit 1 (ND1) primers were used to detect mtDNA or mtRNA, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primers were used to detect nDNA or nRNA. These primer sequences are as follows.

ND1 F: 5′-ACCCCCGATTCCGCTACGACCAAC-3′,

ND1 R: 5′-GGTTTGAGGGGGAATGCTGGAGAT-3′,

GAPDH F: 5′-GGGCAAGGTCATCCCTGAGCTGAA-3′,

GAPDH R: 5′-TCTAGACGGCAGGTCAGGTCCACC-3′.

Calculation of the mtDNA/nDNA, mtRNA/nRNA, and mtRNA/mtDNA ratios

The mtDNA/nDNA, mtRNA/nRNA, and mtRNA/mtDNA ratios were calculated using the equations below. The threshold cycle (Ct) is defined as “the fractional cycle number at which the fluorescence generated by SYBR Green passes above a fixed threshold.”

Ratio of mtDNA/nDNA and ratio of mtRNA/nRNA = 2^(GAPDH Ct- ND1 Ct).

Ratio of mtRNA/mtDNA = (mtRNA/nRNA)/(mtDNA/nDNA).

Measurements of MnSOD, CuZnSOD, Nrf2, Keap1, GPx4 and Catalase gene expressions

The qPCR reactions were performed with the ABI Prism 7000 sequence detection system (Applied Biosystems Foster City, CA, USA) using a QuantiTect SYBR Green PCR Kit (QIAGEN Valencia, CA, USA). GAPDH was used for the control. PCR conditions were as follows: For MnSOD, Nrf2, Keap1, GPx4 amplification, the denature temperature was 95 °C, the denature time was 15 seconds, the annealing and elongation temperature was 60 °C, the annealing and elongation time was 1 minute. For CuZnSDO amplification, the denature temperature was 95 °C, the denature time was 30 seconds, the annealing temperature was 55 °C, the annealing time was 30 seconds, the elongation temperature was 72 °C and the elongation time was 30 seconds. For Catalase amplification, the denature temperature was 95 °C, the denature time was 10 seconds, the annealing temperature was 55 °C, the annealing time was 30 seconds, the elongation temperature was 72 °C and the elongation time was 40 seconds. GAPDH was used as control amplification. These primer sequences are as follows.

MnSOD F: 5′-TTCTGGACAAACCTCAGCCCTAACGGT-3′

MnSOD R: 5′-AACAGATGCAGCCGTCAGCTTCTCCTTAAA-3′

CuZnSOD F: 5′-CATTGCATCATTGGCCGCACACTG-3′

CuZnSOD R: 5′-ACCACAAGCCAAACGACTTCCAGC-3′

Nrf2 F: 5′-GTGGCTGCTCAGAATTGCAGAAAAAGAAAA-3′

Nrf2 R: 5′-TGTTTTTTCAGTAGGTGAAGGCTTTTGTCA-3′

Keap1 F: 5′-CCATGAAGCACCGGCGAAGTGCC-3′

Keap1 R: 5′-GTCTGTATCTGGGTCGTAACACTCCAC-3′

GPx4F: 5′-GAGCCAGGGAGTAACGAAGAGATCAAA-3′

GPx4R: 5′-TCACGCAGATCTTGCTGAACATATCGAATT-3′

Catalase F: 5′-TGACTACGGGAGCCACATCCAGGC-3′

Catalase R: 5′-TCACAGATTTGCCTTCTCCCTTGC-3′

GAPDH F: 5′-GGGCAAGGTCATCCCTGAGCTGAA-3′,

GAPDH R: 5′-TCTAGACGGCAGGTCAGGTCCACC-3′.

Statistical analysis

Statistical analyses were performed using Scheffe’s F test. All p values less than 0.05 were considered to be statistically significant.


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Supplementary Figure 1 X-ray structures of TIRR–53BP1 and NUDT16TI–53BP1.

a, The four TIRR and two 53BP1-Tudor molecules in the asymmetric unit are shown. b, The two NUDT16TI and two 53BP1-Tudor molecules in the asymmetric unit are shown

Supplementary Figure 2 Conformations of TIRR 53BP1-binding loop and corresponding loop region in NUDT16.

Left: Amino acid sequence alignment of TIRR 53BP1-binding loop with corresponding region of NUDT16. Right: Structural overlay of the loop regions in TIRR (blue) and NUDT16 (gray). Highlighted are Pro104 and Arg105 in NUDT16 and key 53BP1-interacting residues Pro105 and Arg107 in TIRR. 53BP1 residues interacting with TIRR Pro105 and Arg107 are also shown.

Supplementary Figure 3 A flexible loop in TIRR (residues 101–107) is highly specific for 53BP1 interaction.

Silver-stained gel (a) and immunoblot (b) of TIRR-FH and TIRR-Loop-FH partner proteins purified from the soluble nuclear extract of U2OS cells for mass spectrometric analysis. TIRR-Loop-FH corresponds to TIRR-FH mutated to harbor the loop region of NUDT16 (see Supplementary Fig. 2). Mass spectrometry data are in Supplementary Table 1

Supplementary Figure 4 Class switch recombination in stimulated B cells.

Shown are representative flow cytometry plots of the data presented in Fig. 5h

Supplementary Figure 5 Comparison of the X-ray structures of TIRR–53BP1 and NUDT16TI–53BP1.

a, Overlay of the TIRR–53BP1 and NUDT16TI–53BP1 structures with TIRR shown in blue, NUDT16TI in light blue, and 53BP1 in orange. The C-terminal α-helices in TIRR homodimer (shown in gray) do not exist in NUDT16TI. b, Details of the TIRR–53BP1 and NUDT16TI–53BP1 binding interfaces illustrating the remarkable similarity between the two complexes. Same color-coding as in a.


Enzymatic Hydrolysis and Simultaneous Saccharification and Fermentation of Soybean Processing Intermediates for the Production of Ethanol and Concentration of Protein and Lipids

Carbohydrates in soybeans are generally undesirable due to their low digestibility and because they “dilute” more valuable components (proteins, lipids). To remove these carbohydrates and raise the titer of more valuable components, ethanol production was investigated. Commercial enzymes (Novozyme cellulase, β-glucosidase, and pectinase) were added to ground soybeans (SB), soybean meal (SBM), soybean hulls (SH), and soybean white flakes (WF) at a 10% solids loading rate to quantify hydrolyzed glucan. Saccharification resulted in glucan reductions of 28%, 45%, 76%, and 80% (SBM, SB, SH, WF, resp.). Simultaneous saccharification and fermentation (SSF) trials were conducted at 5%, 10%, 15%, and 20% solids loading with Saccharomyces cerevisiae NRRL Y-2034 and Scheffersomyces stipitis NRRL Y-7124, with protein, fiber, and lipids analyzed at SSF 10% solids and saccharification trials. S. cerevisiae and S. stipitis produced

2.5–8.6 g/L ethanol, respectively, on SB, SBM, and WF over all solid loading rates. SH resulted in higher ethanol titers for both S. cerevisiae (

9–23 g/L) and S. stipitis (

9.5–14.5 g/L). Protein concentrations decreased by 2.5–10% for the SB, SBM, and WF, but increased by 53%–55% in SH. Oil concentrations increased by

1. Introduction

Soybeans are one of the most valuable crops in the world due to their high oil and protein content, which provides for a wide variety of uses. Soybean oil is used as a food and feed ingredient as well as in cosmetics [1–4] and biodiesel production [5]. Soybean protein is highly digestible and has been used in livestock and aquaculture feeds, along with many human foods [6–10]. Soybean protein supplements are promoted in human diets due to their many health benefits [9, 11–15].

In contrast to the oil and proteins, carbohydrates found in soybeans, (

10% dry weight), are largely undesirable due to their low digestibility [16]. Stachyose and raffinose, two of the primary carbohydrates in soybeans, are indigestible by humans and other monogastrics but can be fermented by natural flora in the intestinal tract, causing discomforting gas buildup [17–19]. Stachyose and raffinose may also decrease the digestibility of foods which contain them [17]. The presence of these carbohydrates also effectively dilutes the concentration of the protein and oil in soybeans.

In many countries, soybeans are “crushed” and then extracted with hexane or other solvents to separate soybean oil from the solids (i.e., soybean meal, SBM). SBM has a protein content of about 45% and is widely used as a livestock feed. Soybeans can be further processed via ethanol extraction to remove carbohydrates, resulting in soy protein concentrate (SPC) that contains at least 65% protein [18]. This is an expensive process and the removed sugars have little use, but the SPC is much more digestible and commands a price 2–2.5 times that of SBM.

As an alternative to ethanol washing, we evaluated saccharification and bioconversion of soybean carbohydrates to ethanol. This would create an additional product to help offset processing costs, while making use of an underutilized material (soybean carbohydrates). Commercial hydrolytic enzymes and yeast were tested on soybeans and three fractions from the soybeanoil extraction facility. We hypothesized that much of the protein or lipids used by yeast as nutrients would be left in the final solids as yeast cell mass. Moreover, we expected an increase in protein content due to the conversion of carbohydrates into cell protein.

2. Materials and Methods

2.1. Substrates

Substrates tested included whole soybeans, soybean hulls, white flakes, and defatted soybean meal. These were obtained as a gift from South Dakota Soybean Processors, Volga, SD, USA. Figure 1 shows a simplified process flow diagram of organic-solvent soybean processing to denote the source of the substrates. The substrates were ground using a Wiley Mill (2 mm) screen and stored at room temperature. These substrates were subjected to proximate analysis by Olson Agricultural Analytical Laboratories at South Dakota State University in Brookings, SD, USA and the results are shown in Table 1.


2.2. Enzymes

Enzymes were obtained as a gift from Novozymes (Franklinton, NC, USA). NS 50013 (Celluclast 1.5 L) is a cellulase cocktail with an activity of 70 FPU/g. NS 50010 (Novozyme 188) is a β-glucosidase with an activity of 250 CBU/g. NS 22016 is a pectinase cocktail with an activity of 3800 U/mL. Enzymes were stored at 4°C prior to use.

2.3. Yeast

Saccharomyces cerevisiae NRRL Y-2034 and Scheffersomyces stipitis NRRL Y-7124 were obtained from the USDA ARS Culture Collection (Peoria, IL, USA). For short-term maintenance, cultures were grown on Potato Dextrose Agar (PDA) plates and slants for 72 h at 35°C and then stored at 4°C, with subculturing of the organisms every 4 weeks. Lyophilization in a 20% sucrose solution was used for long-term storage.

Inoculum for all trials was prepared by aseptically inoculating sterile 5% glucose, 0.5% yeast extract broth (100 mL in a 250 mL Erlenmeyer flasks) with a 1% (v/v) aliquot for S. cerevisiae, or 5% (v/v) for S. stipitis, from broth seed cultures stored at 4°C. Flasks for inoculum were incubated for 24 h at 35°C in a 250 rpm rotary shaker. Broth seed cultures were grown for 24 h at 250 rpm before refrigeration and used within 60 days to inoculate flasks for inoculum.

2.4. Buffers and Antibiotics

Saccharification and SSF trials were conducted in a sterile 0.1 M sodium citrate buffer with the pH adjusted to 4.8 using concentrated H2SO4. A stock solution of 10 mg/mL tetracycline HCl (70% ethanol and filter-sterilized) was prepared and stored at −20°C, from which 0.4 mL/100 mL of total trial volume was used to prevent bacterial contamination. A stock solution of 10 mg/mL cycloheximide (filter-sterilized) was prepared and stored at 4°C, from which 0.3 mL/100 mL of total trial volume was used for contamination control in saccharification trials only.

2.5. Saccharification of Soybean Fractions

Saccharification trials were conducted by mixing 15 g of ground substrate with 75 mL of sterile 0.1 M sodium citrate buffer, along with tetracycline and cycloheximide solutions in 250 mL Erlenmeyer flasks fitted with rubber stoppers. The pH of the solutions was adjusted to 5.0 using concentrated H2SO4 or 12 M NaOH. The stoppers were pierced with 21 gauge syringe needles and Whatman 0.2 μm syringe filters. Enzyme dosages per gram of glucan included 23.2 FPU of NS 50013, 41 CBU of NS50010, and 500 U NS 22016. Table 2 provides a summary of glucan levels found in the literature for each soybean fraction, and these were averaged to calculate enzyme dosage. Table 3 shows the volume of enzymes used for each substrate. Sterile-deionized water was added to each flask to bring the total volume to 150 mL, resulting in a solid loading rate of 10%. Saccharification trials were run for 96 h in a 50°C reciprocating shaker set at 250 rpm. Flasks lacking enzymes were used as controls to determine the type and amount of carbohydrates that would be released by solubilization.

2.6. Simultaneous Saccharification and Fermentation of Soybean Fractions

Ground substrates (15, 30, 45, or 60 g) were mixed with 150 mL of sterile 0.1 M sodium citrate buffer, along with an appropriate amount of tetracycline solution in 500 mL Erlenmeyer flasks fitted with rubber stoppers that were pierced with 21 gauge syringe needles and attached to Whatman 0.2 μm syringe filters. The pH was adjusted to 5.0 using concentrated H2SO4 or 12 M NaOH. The substrates were not autoclaved in an effort to preserve protein and carbohydrate integrity by preventing the Maillard reaction [25] as well as retaining the most likely parameters for industrial applications. Enzyme dosages per gram of glucan included 23.2 FPU of NS 50013, 41 CBU of NS 50010, and 500 U of NS 22016. Table 2 shows the amount of substrate, fiber, and enzymes used at the 5% loading rate for each substrate. Amounts of these components increased proportionally at the higher solid loading levels. Sterile-deionized water was added to bring the total volume to 297 mL, and then 3 mL of a 24 h culture of either S. cerevisiae or S. stipitis was added. Flasks were incubated for 96 h in a 35°C reciprocating shaker set at 250 rpm. Control flasks without enzymes were also included to assess ethanol production from the free carbohydrates released from the substrates. These controls were prepared in the same manner as described above, except that the volumes of enzymes were replaced with sterile-deionized water.

2.7. Analytical Methods

Samples (5–10 mL) were drawn aseptically from the flasks at 0, 2, 4, 6, 12, 24, 48, 72, and 96 h. Samples were boiled for 5 minutes to inactivate enzymes and then centrifuged at 2400 ×g for 10 min. After freezing for 24 h at −20°C, samples were thawed, centrifuged again at 13,000 rpm for 15 min, and the supernatant was filtered through 0.2 μm syringe filter into HPLC autosampler vials.

Carbohydrates were analyzed using a Waters 1200 HPLC with a Waters Sugar-pak I column and refractive index detector. The mobile phase for the Sugar-pak I column was 0.0001 M calcium EDTA flowing at a rate of 0.5 mL/min, with the column at 65°C. Ethanol concentrations were determined using a Waters 717 HPLC with an Aminex HPX-87H column and Waters 2410 refractive index detector (RID). The mobile phase was 0.005 M H2SO4 flowing at a rate of 0.6 mL/min, with the column at 65°C.

At the end of saccharification and SSF trials, in the 10% solid loading rate experiments, the slurries from all replicates of a trial were combined and evaporated to dryness in an 80°C oven for 96 h. A proximate analysis was performed on the solids by Olson Agricultural Analytical Laboratory Services (South Dakota State University, Brookings, SD, USA).

2.8. Data Analysis

Saccharification trials were done in replicates of six, while the SSF trials were done in triplicate. Parameters analyzed included maximum ethanol concentration, ethanol productivity, and residual carbohydrates. The percent difference of the fiber, protein, and lipid content when compared to the original substrate was calculated using the formulas listed below. Residual carbohydrates were corrected by subtracting additional carbohydrate results that resulted from denatured enzymes or buffer. Graphs and calculations were made in Microsoft Excel 2007. (i) Ethanol Productivity (g/L/h) = (Net Maximum Ethanol Concentration)/Time. (ii) Component Percent Difference (%) = ((% of dried slurry after trial) − (% of original substrate))/% of original substrate.

3. Results and Discussion

3.1. Saccharification of Soybean Fractions

Table 4 shows the composition of the four soybean substrates following 96 h saccharification. Trials (six replicates) were conducted at the 10% solid loading rate, both with and without enzymes. Soluble carbohydrate levels in the saccharified broth were determined via HPLC for each individual replication. Following saccharification, solids from the six replications of each treatment were combined, dried, and analyzed for fiber, protein, and lipid levels. The percentage difference for the fiber, protein, and lipid concentrations, compared to the substrate before saccharification, was calculated.

As expected, the presence of enzymes resulted in higher soluble carbohydrate levels for each of the substrates, compared to control trials lacking enzymes. This difference was statistically significant for all substrates except white flakes and was also correlated with the reduction in fiber content. Whole beans contained the lowest concentration of fiber, due to the presence of both lipids and protein. Consequently, soluble carbohydrates were the lowest and only a moderate reduction in percent fiber was observed. Enzymatic saccharification efficiency in raw beans may have been reduced by the lack of any pretreatment effect that the typical soy processing operation provides. On the other hand, hulls contained the highest level of fiber and therefore responded most significantly to enzymatic hydrolysis, yielding the highest level of carbohydrates and greatest percent reduction in fiber.

In the soybean crushing process, after oil extraction the solids are referred to as white flake. This material is then heated to drive off any residual hexane and inactivate certain antinutritional factors. Low levels of hulls then may be added to white flake to reduce the protein content to

45%. This material is then called soybean meal. Thus, white flake and soybean meal are relatively similar in composition and we anticipated similar results upon saccharification. As can be seen in Table 4, soybean meal resulted in higher soluble carbohydrates and a greater effect of enzyme addition. Perhaps this was due to the additional heat treatment and/or presence of some hulls.

Protein and lipid levels in the samples were not expected to change significantly, since only small amounts of enzymes, buffers, and other components were added and the total solids were recovered. The only significant change expected was the conversion of fiber to soluble sugars as described above. Changes in relative protein levels varied from −14% to +11% and showed no significant trends. Lipid levels in whole beans increased from 4.5% to 12.9%, again likely due to greater solubilization during the saccharification process. Lipid levels in the other materials were very low, and therefore slight variability in values resulted in large percent changes.

3.2. Soybean Substrate SSF

Each substrate was subjected to SSF treatment using four different concentrations of substrate (5%, 10%, 15%, or 20% solid loading rate (SLR)) as well as either S. cerevisiae or S. stipitis. Enzyme dosages were normalized based on glucan levels and control trials lacking enzymes were also performed. Carbohydrate and ethanol titers were monitored throughout each 96 h SSF. After SSF, the replicates from the 10% solid loading rate treatments were combined and dehydrated for fiber, protein, and lipid analysis by Olson Agricultural Analytical Laboratory Services.

Figure 2 shows that the maximum ethanol titer of both yeasts increased as the SLR of soybeans was increased from 5% to 20%, as was expected. In most treatments, S. cerevisiae produced more ethanol than S. stipitis (maxima of 12.357 g/L ± 5.213 for S. cerevisiae with enzymes, 7.726 g/L ± 1.167 for S. stipitis). However, the difference was only significant in the 15% SLR trial with enzymes and the 20% SLR trial without enzymes. The presence of enzymes enhanced ethanol levels, but was only statistically significant in the 5% SLR trials. The high degree of variability in the 15% and 20% SLR trials was likely due to the high viscosity of these trials, which reduced mixing efficiency.


Maximum ethanol titer from soybeans after 96 h SSF or fermentation 1 . 1 Error bars represent one standard deviation.

Figure 3 shows the corresponding ethanol productivities for the soybean SSF or fermentation trials, which were calculated at the time of maximum ethanol concentration. As expected, ethanol productivities also increased as SLRs increased from 5% to 20%. In most comparisons, S. cerevisiae had significantly higher productivities compared to S. stipitis. Ethanol productivities were actually higher in many of the enzyme-free trials (maxima of 0.397 g/L/h ± 0.127 for S. cerevisiae and 0.134 g/L/h ± 0.107 for S. stipitis, both 15% SLR without enzymes), suggesting that enzymatic hydrolysis of the nonpretreated soybean was the rate limiting factor.


Ethanol productivity from soybeans after 96 h SSF or fermentation 1 . 1 Error bars represent one standard deviation.

Figure 4 shows the total residual carbohydrate levels after 96 h SSF or fermentation. The levels of residual carbohydrates increased as the SLR increased, reflecting an accumulation of stachyose, raffinose, or partial hydrolysis products of the oligosaccharides. This was due to the inability of either yeast to fully catabolize the oligosaccharides [26]. S. cerevisiae can hydrolyze the fructose residue from both stachyose and raffinose by use of invertase [27, 28], but cannot hydrolyze the other bonds. S. stipitis does not produce invertase and cannot catabolize either oligosaccharide. In most treatments, total residual carbohydrate levels were similar between yeasts however, at the 10% and 20% SLR trials with enzymes, S. stipitis accumulated significantly higher carbohydrate levels than S. cerevisiae. Also expected were the higher carbohydrate levels in enzyme-hydrolyzed trials compared to enzyme-free trials. The highest carbohydrate concentration was 20% SLR with enzymes and S. stipitis (18.76 g/L ± 5.501), and the lowest was 5% SLR without enzymes with S. cerevisiae (1.96 g/L ± 0.661).


Residual carbohydrates from soybeans after 96 h SSF or fermentation 1 . 1 Error bars represent one standard deviation.

Figures 5–7 show the maximum ethanol titers, ethanol productivities, and residual carbohydrate levels for SSF and fermentation trials with hulls. Since the hulls contain primarily fiber, and lower levels of oligosaccharides than the other soybean fractions, we anticipated that ethanol production would be enhanced. Figure 5 shows a statistically significant difference in ethanol production between SSF trials with versus without enzymes as well as increased ethanol production as the SLR increased (except between 15% and 20% SLR). S. cerevisiae outperformed S. stipitis at the higher SLRs, perhaps due to increased ethanol tolerance. Maximum concentrations obtained were 23.177 g/L ± 10.148 for S. cerevisiae and 14.501 g/L ± 6.748 for S. stipitis. As with the beans, the hulls were highly viscous at the higher SLRs, making proper mixing of the slurry very difficult, adding to the variability of the trials.


IHC Troubleshooting Guide

The following image provides an example of IHC staining.

Immunohistochemistry of formalin-fixed paraffin-embedded (FFPE) cancer tissue. Analysis was performed to compare Connexin 43 membrane staining in FFPE sections of human lung adenocarcinoma (right) compared to a negative control without primary antibody (left). To expose target proteins, heat-induced epitope retrieval (HIER) was performed using 10 mM sodium citrate (pH 6.0), followed by heating in a microwave for 8 to 15 minutes. After HIER, tissues were blocked in 3% H2O2 in methanol for 15 minutes at room temperature, washed with distilled H2O and PBS, and then probed overnight at 4°C in a humid environment with an Invitrogen Connexin 43 monoclonal antibody (Cat. # 13-8300), diluted 1:20 in PBS/3% (w/v) BSA. Tissues were washed extensively in PBS buffer containing 0.05% (v/v) Tween-20 (PBST). Detection was performed using an HRP-conjugated secondary antibody followed by chromogenic detection using DAB as the substrate. The sections were counterstained with hematoxylin and dehydrated with ethanol and xylene prior to mounting.

The following points are provided to help identify the cause of high background staining, which results in a poor signal-to-noise ratio. See also the additional notes sections at the bottom of this page for more information.

Cause: Endogenous enzymes

Incubate a test tissue sample with the detection substrate alone for a length of time equal to that of the antibody incubation. A strong background signal suggests interference from endogenous peroxidases or phosphatases.

  • Solution: Quench endogenous peroxidases with 3% H2O2 in methanol or water or use a commercial kit, such as Thermo Scientific Peroxidase Suppressor.

Endogenous phosphatases can be inhibited with the endogenous alkaline phosphatase inhibitor, levamisole.

Cause: Endogenous biotin or lectins

High background can occur when endogenous biotin is not blocked prior to adding the avidin–biotin–enzyme complex.

If the ABC complex is made with avidin, the highly-glycosylated protein can bind to lectins in the tissue sample.

  • Solution: Block endogenous lectins with 0.2 M alpha-methyl mannoside in dilution buffer. Alternatively, use streptavidin or Thermo Scientific NeutrAvidin Protein instead of avidin, because both are not glycosylated and won't bind to lectins.

Cause: Secondary antibody cross-reactivity or nonspecific binding

The secondary antibody may show a strong or moderate affinity for identical or similar epitopes on non-target antigens.

  • Solution: If normal serum from the source species for the secondary antibody is used to block the tissue, then increase the serum concentration to as high as 10% (v/v), if necessary. If you are blocking with another reagent (BSA, nonfat dry milk), then add 2% (v/v) or more normal serum from the source species for the secondary antibody. Alternatively, reduce the concentration of the biotinylated secondary antibody.

Egg white, which contains avidin, was often used to coat slides, dilute antibodies or block tissue samples because it is a readily available and inexpensive source of carrier proteins. It is used very rarely nowadays.

  • Solution: Avoid using egg whites to prevent egg white–based avidin from binding biotinylated secondary antibody during IHC staining. Synthetic tissue adhesives as well as avidin-free antibody diluents and blocking buffers are readily available.

Cause: Issues with the primary antibody

Nonspecific interactions between the primary antibody and non-target epitopes in the tissue sample occur regularly during incubation but at a level that does not influence background staining. A high primary antibody concentration will increase these interactions and thus increase nonspecific binding and background staining.

The primary antibody may also show a strong or moderate affinity for identical or similar epitopes on non-target antigens.

  • Solution: Increase the blocking buffer composition and/or concentration, or use a different primary antibody.

The primary antibody diluent may contain little or no NaCl, which helps to reduce ionic interactions.

  • Solution: Add NaCl to the blocking buffer/antibody diluent so that the final concentration is between 0.15 M and 0.6 M NaCl. The best NaCl concentration to use will have to be determined empirically.

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See also the additional notes sections at the bottom of this page for more information.

Cause: Enzyme–substrate reactivity

Even when the tissue sample is properly prepared and labeled, the enzyme–substrate reaction must occur for the chromogenic precipitate to form. Deionized water can sometimes contain peroxidase inhibitors that can significantly impair enzyme activity. Additionally, buffers containing sodium azide should not be used in the presence of HRP. Finally the pH of the substrate buffer must be appropriate for that specific substrate.

A simple test to verify that the enzyme and substrate are reacting properly is to place a drop of the enzyme onto a piece of nitrocellulose and then immediately dip it into the prepared substrate. If the enzyme and substrate are reacting properly, a colored spot should form on the nitrocellulose.

Solution: Change the enzyme diluent and/or prepare substrate at the proper pH and repeat the test.

Cause: Primary antibody potency

Primary antibodies generally lose affinity for the target antigen over time, either due to protein degradation or denaturation caused by long-term storage, microbial contamination, changes in pH or harsh treatments (e.g., freeze/thaw cycles).

Test the primary antibody for potency by staining tissue samples known to contain the target antigen with various concentrations of the primary antibody do the test concurrently with the test sample. If the positive control is not positive for the target antigen at all, then this suggests that the primary antibody has lost potency. In fact, it is good laboratory practice to always run a positive control sample through your staining protocol along with the experimental samples.

Solution: Ensure that the antibody diluent pH is within the specified range for optimum antibody binding (7.0 to 8.2) and that the antibody is stored according to the manufacturer's instructions. To prevent contamination of your antibody solutions, wear gloves when dispensing antibodies, and use sterile pipette tips, if appropriate. Even if you store your antibodies in a refrigerator, always divide them into separate small aliquots. This prevents contamination or loss of the whole vial of antibody if a problem arises.

Cause: Secondary antibody inhibition

While high concentrations of the secondary antibody can increase background staining, extremely high concentrations can have the opposite effect and reduce antigen detection.

To test if the secondary antibody concentration is inhibitory, stain positive control samples using decreasing concentrations of the secondary antibody. An increase in signal as the concentration decreases suggests that antibody concentration is too high.

Solution: Reduce the concentration of the secondary antibody.

If the diluent and/or blocking solution contains antigen-neutralizing antibodies, such as those found in serum, then the antibodies will block secondary antibody binding.

Solution: Remove the neutralizing antibodies or change to a different diluent and/or blocking solution.


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Watch the video: Make Sodium ethoxide (August 2022).