What Is the Ph of 0025 M Solution of Thymol Blue

Volume 3

Kyung J. Kwon-Chung , in The Yeasts (Fifth Edition), 2011

Key to Species

1.	a. Yeast state is present..........2 b. Yeast state is not present..........3
2(1).	a. Basidiospores are finely roughened, subglobose to ellipsoid, d -proline is not assimilated, reaction on canavanine–glycine–brom thymol blue (CGB) agar is negative in 2–5 days.......... F. neoformans: b. Basidiospores are smooth, bacilliform with or without a curvature, d-proline is assimilated, CGB agar reaction is positive..........F. bacillispora:	p. 1449
		p. 1445
3(1).	a. Basidiospores are obpyriform to pentagonal, up to 4   μm in diameter..........F. depauperata: b. Basidiospores are globose to subglobose, 4–7   μm in diameter..........F. lutea:	p. 1447
		p. 1448

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780444521491001142

Cinnarizine

Nadia G. Haress , in Profiles of Drug Substances, Excipients and Related Methodology, 2015

4.2.2 Spectrophotometric Methods

4.2.2.1 Ultraviolet Spectrometry

Hanmin and Xiuquan [13] reported the determination of cinnarizine in tablets by ultraviolet spectrophotometry based on the measurement of absorbance at 250   nm. Standard plots of absorbances and the concentrations of cinnarizine were linear for 5–25   μg/mL of cinnarizine. Recoveries were 93.9–98.9%.

Saleh and Askal [14] described a spectrophotometric method for the determination of cinnarizine in capsules and tablets based on the charge-transfer complex formation between cinnarizine as n-donor and either iodine as δ-acceptor or 2,3-dichloro-5,6-dicyano-p-benzoquinone (DDQ) as n-acceptor. Ranges for obedience of absorbance at 295 and 460   nm to Beer's law for cinnarizine with iodine and DDQ were 1–6 and 10–8   μg/mL, respectively. Recoveries of cinnarizine were ~   100% and standard deviations were 0.72–1.15%.

Abdine et al. [15] developed a direct, extraction-free spectrophotometric method for the determination of cinnarizine in pharmaceutical preparations. The method was based on ion-pair formation between cinnarizine and three acidic (sulfonephthalein) dyes, namely bromocresol green, bromocresol purple, and bromocresol blue which induced an instantaneous bathochromic shift of the maximum in the drug spectrum. Conformity to Beer's law enabled the assay of dosage forms of cinnarizine. Compared with a reference method, the results obtained were of equal accuracy and precision.

Elazazy et al. [16] used a simple, rapid, sensitive, and accurate spectrophotometric method for the determination of cinnarizine, famotidine, and metoclopramide hydrochloride in pure form and in pharmaceutical formulations. The spectral method is based on the reaction between dichlorophenol indophenol and the cited drugs to give bluish violet radical ions exhibiting maximum absorption at 650, 642, and 654   nm for cinnarizine, famotidine, and metoclopramide, respectively, with molar absorptivities 2.421   ×   10³, 4.313   ×   10³, and 2.112   ×   10⁴  L/mol/cm for cinnarizine, famotidine, and metoclopramide, respectively, and Sandell's sensitivities 6.569   ×   10^{−   3}, 1.278   ×   10^{−   3}, and 6.646   ×   10^{−   3}  μg/cm².

Devagondanahalli et al. [17] described two simple, rapid, and sensitive extractive spectrophotometric methods for the assay of cinnarizine in pure and pharmaceutical formulations. The spectrophotometric methods depend on the formation of chloroform-soluble ion-association complexes of cinnarizine with thymol blue (TB) and with cresol red in sodium acetate–acetic acid buffer of pH 3.6 for TB and in potassium chloride–hydrochloric acid buffer at pH 1.6 for cinnarizine with absorption maxima at 405 and 403  nm for TB and cinnarizine, respectively. Reaction conditions were optimized to obtain the maximum color intensity. The systems obeyed Beer's law in the range of 0.6–15.8 and 0.8–16.6   μg/mL for TB and cinnarizine, respectively. Various analytical parameters have been evaluated and the results have been validated by statistical data.

Tarkase et al. [18] reported the development and validation of spectrophotometric method for simultaneous estimation of cinnarizine and domperidone maleate in pure and tablet dosage form. The spectral method depends on simultaneous equation method at two selected wavelength 254 and 284   nm, respectively, and also on absorbance ratio method at two selected wavelengths 274   nm (isoabsorptive point) and 254   nm (λ _max of cinnarizine). The linearity was obtained in the concentration range of 5–20 and 5–20   μg/mL for cinnarizine and domperidone maleate, respectively. These methods are accurate, precise, reproducible, and economical, and the results have been validated statistically and by recovery studies.

Abdelrahman [19] described a simultaneous determination of cinnarizine and domperidone in a binary mixture by using area under the curve and dual wavelength spectrophotometric methods. In area under the curve method, mixture solutions in the wavelength ranges 241–258 and 280–292   nm were selected for the determination of cinnarizine and domperidone, and by applying Cramer's rule the concentration of each drug was obtained. In dual wavelength method, two wavelengths were selected for each drug in a way so that the difference in absorbance is zero for another drug. Domperidone shows equal absorbance at 240.2 and 273.2   nm, where the differences in absorbance were measured for the determination of cinnarizine. Similarly, differences in absorbance at 230.8 and 239.2   nm were measured for determination of domperidone. The spectral methods were applied for the determination of cinnarizine and domperidone over the concentration ranges of 2–20 and 2–22   μg/mL, respectively. Both methods were found to be simple, accurate, sensitive, precise, and inexpensive which could be used in routine and quality control analysis of the cited drugs in pharmaceutical formulations containing them.

Issa et al. [20] reported the determination of cinnarizine in pure and in its pharmaceutical dosage forms by spectrophotometric methods carried out to investigate the charge-transfer complex formation between cinnarizine and dipicrylamine (DPA) or 2,6-dinitrophenol (DNP). The colored products were quantified spectrophotometrically at 430 and 440   nm for cinnarizine complexes with DPA in a mixture of 15% dioxane in dichloroethane and 15% ethyl acetate in chloroform, respectively. On the other hand, cinnarizine complexes formed with DNP in acetonitrile and in a mixture of 30% dichloroethane in ethyl alcohol were quantified at 460 and 430   nm, respectively. Beer's law was obeyed in the concentration range of 1–36.8   μg/mL. These methods utilize a single-step reaction and have the advantages of being simple, accurate, sensitive, rapid, and suitable for routine analysis in control laboratories.

4.2.2.2 Spectrofluorimetric Method

Walash et al. [21] suggested the use of second-derivative synchronous fluorimetric method for the determination of cinnarizine and domperidone in different pharmaceutical formulations. The fluorimetric method is based upon measurement of the native fluorescence of these drugs at λ _max 315 and 324   nm for cinnarizine and domperidone, respectively, after excitation at 280   nm. The synchronous fluorescence spectra of cinnarizine with domperidone were recorded using the optimum Δλ 80   nm in aqueous methanol (50% v/v). The produced fluorescence-concentration plots were rectilinear obeying Beer's law in the concentration range of 0.1–1.3 and 0.1–3   μg/mL for cinnarizine and domperidone, respectively, with lower detection limits of 0.017 and 5.77   ×   10^{−   3}  μg/mL and quantification limits of 0.058 and 0.02   μg/mL for cinnarizine and domperidone, respectively. This simple, rapid, and highly sensitive method was successfully applied for the determination of cinnarizine in biological fluids.

4.2.2.3 Chemiluminescence Method

Townsend et al. [22] reported a flow-injection chemiluminescence method for the determination of cinnarizine by using the chemiluminescence of permanganate system in the presence of polyphosphoric acid, ethanol, and Tween 60. Five hundred-microliter samples were injected and the sample throughput was 130   h^{−   1}. Preliminary experiments identified Tween 60 as the surfactant of choice, improving the detection limit of cinnarizine system 20-fold. Optimum chemiluminescence signals were obtained using 7.5   ×   10^{−   4}  mol/L potassium permanganate in 0.02   mol/L polyphosphoric acid as the oxidant stream and a carrier stream of 10% (v/v) of ethanol in aqueous 1.5   ×   10^{−   3}  mol/L Tween 60 with a total flow rate of 7.6   mL/min. The calibration curve was linear from 0.5 to 6   μg/mL with recoveries of 98.4–100.2%.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/S1871512515000023

Analysis of fermented foods

Osman Erkmen , in Microbiological Analysis of Foods and Food Processing Environments, 2022

33.2.1 Equipment, materials, and media

Equipment. Blender (or Stomacher), capped jar, cover glass, Erlenmeyer flask, filter paper (0.45   μm), gasometer, hemocytometer counting chamber, incubator, microscope glass slide, microscope, Petri dishes, pH meter, pipettes (1 and 10   mL) in pipette box, pipette discarding pad (containing 10% bleach solution), platinum loop (3   mm diameter), test tubes, thermostatic shaking water bath, and vortex.

Materials. BaCl₂ , calcium citrate solution, chloroform, creatin, erythrosine stain (0.02%), Gram staining reagents, HCl, methyl thymol blue (MTB) solution (0.1%), NaOH (0.1, 0.5 and 1.0N), NaOH (40%), 0.1% peptone water, phenolphthalein indicator (0.5%), potassium chromate solution (5%), quarter-strength Ringer's (QSR) solution, silver nitrate solution (0.171N; 29.063  g   L⁻¹), sterile distilled water, trichloroacetic acid solution (4%), and xylen.

Media. 2.0% Agar, APT agar, bromocresol green ethanol (BGE) agar, chopped liver (CL) broth, dextrose sorbitol mannitol (DSM) agar containing bromocresol, Elliker agar, glucose yeast extract calcium carbonate (GEYC) broth supplemented with 100   mg pimaricin per L, HHD agar containing fructose, Lee's agar, litmus milk, M17 agar, de Man-Rogosa and Sharpe (MRS) agar, MRS broth, MRS broth containing 1%, 1.5%, and 2% bile salt, MRS broth (with pH 2.0, 3.0, 4.0, and 5.0), MRS supplemented with cysteine hydrochloride (mMRS) containing novobiocin and van comycin, neutral red chalk lactose (NRCL) agar, nitrate peptone water, Rogosa agar, skim milk (10%), yeast extract skim milk (YESM) agar, yeast extract glucose (YEG) broth, YEG broth containing acetic acid from 0.2% to 7.0%, YEG broth containing ethanol from 6% to 15%, yeast extract glucose lemco (YEGL) bromocresol purple agar, and yeast extract peptone (GYEP) agar.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780323916516000239

Marine Enzymes and Specialized Metabolism - Part B

Ron Wever , ... Rokus Renirie , in Methods in Enzymology, 2018

2.4 Activity Assays for IPO and BPO Activity

Vanadium-dependent iodo- and BPOs are unable to oxidize directly the classical heme peroxidase substrates such as guaiacol, o-dianisidine, and benzidine, and thus other assays should be used for assessing enzymatic activity. However, to detect activity on polyacrylamide gels or for localization of the peroxidase microscopically in cross sections of algal part, o-dianisidine may be used. The gels or cross sections are first incubated in a buffered solution containing 100   mM KBr or 100   mM KI, and 10   mM o-dianisidine, and the staining is started by addition of 2   mM H₂O₂. The formed HOBr or HOI will oxidize the organic dye resulting in polymerization and formation of a brown precipitate. Please note that o-dianisidine is carcinogenic and the solutions should be handled carefully.

2.4.1 IPO Activity

Classically (Björkstén, 1968) the IPO activity of peroxidases was assessed by the spectrophotometric detection of triiodide formed during the enzymatic reaction according to:

(1) ${\mathrm{H}}_2{\mathrm{O}}_2+{\mathrm{I}}^{-}\to \mathrm{HOI}+{\mathrm{H}}_2\mathrm{O}$

(2) $\mathrm{HOI}+{\mathrm{I}}^{-}+{\mathrm{H}}^{+}\to {\mathrm{I}}_2+{\mathrm{H}}_2\mathrm{O}$

(3) ${\mathrm{I}}_2+{\mathrm{I}}^{-}\leftarrow \to {{\mathrm{I}}_3}^{-}$

The VIPOs oxidize iodide to HOI (Eq. 1), and HOI will reacts further to I₂ (Eq. 2). I₂ will form a reversible and weak complex with I⁻ according to Eq. (3) to form ${{\mathrm{I}}_3}^{-}$ which absorbs at 350   nm (ɛ  =   25   mM ^{−   1}  cm^{−   1}). The equilibrium constant value of 830 M ^{−   1} in Eq. (3) (Eigen & Kustin, 1962) is high, and IPO activity can only be measured quantitatively at iodide concentrations higher than 20   mM. Due to the requirement of high I⁻ concentrations to detect I₂ formation as ${{\mathrm{I}}_3}^{-}$ , the K _m values determined by this assay will be stacked in the mM region and the published values do not reflect a true K _m for iodide. Further a reaction occurs between HOI and H₂O₂ at pH values >   7.0 yielding singlet oxygen (Verhaeghe, Buisson, et al., 2008), and thus reported values in the literature for K _m and V _max should be treated with caution. In fact, the triiodide assay should only be used for qualitative applications (Verhaeghe, Buisson, et al., 2008). An alternative and quantitative spectrophotometric assay for the determination of IPO activities of VHPO has been developed (Verhaeghe, Buisson, et al., 2008 ) based on the iodination of thymol blue. Thymol blue has a high reactivity toward HOI, and the steady-state kinetic of the oxidation of iodide by VBPO from A. nodosum has been studied. This method illustrated in Fig. 2 is better than the standard ${{\mathrm{I}}_3}^{-}$ assay but, unfortunately, can only be used in the pH range 7–8.

Fig. 2. Bromination or iodination of thymol blue. At pH 8 there is marked color change due a shift in the pK _a upon halogenation of the dye, the dihalogenated compound has a deep blue color with an absorbance maximum at 620   nm. The diiodo-thymol sulfonphtalein has an absorbance coefficient of 40.3   mM ^{−   1}  cm^{−   1} and the absorbance coefficient of the dibrominated compound is 37.2   mM ^{−   1}  cm^{−   1}.

The marked color change from yellow to deep blue in the assay is restricted to pH values above 7 due to the change in pK _a of thymol blue (8.9) to the pK _a (7.3) of the formed iodinated product. Thus, at lower pH kinetic parameters of the VIPOs or their pH optima cannot be assessed directly by this method.

2.4.2 BPO Activity

The activity of the bromo- and CPOs is routinely determined using scavengers reacting with the reactive intermediates HOCl or HOBr and that result in marked changes in absorbance.

2.4.2.1 Phenol Red Assay

A useful assay is the halogenation of the water-soluble version of dye phenol red (Fig. 3) that may be used only for qualitative purpose since the mono-, di-, tri-, and tetrabrominated bromophenol are formed sequentially (De Boer et al., 1987), and these compounds differ in their absorption maxima. In addition, the reactivity of the phenol red is fairly low and competition with other scavengers may occur.

Fig. 3. Bromination of phenol red used to detect halogenating activity qualitatively. Owing to bromination, a marked color change occurs from yellow (at acidic to neutral pH) to deep blue or purple, which can easily be detected by eye.

However, the marked color change allows visual screening of haloperoxidase activity of large numbers of samples or screening of seaweeds on the presence of haloperoxidases on the site of collection (Suthiphongchai et al., 2008). It may also be used to rapidly detect and qualitatively assess activity of fractions during ammonium sulfate fractionations and of eluents of columns during the purification procedures (see Sections 3.1 and 3.2). Similarly, it may be used in kinetic laboratory experiments for undergraduate laboratory exercises (Jervis, Jervis, & Giovannelli, 2005; Jittam et al., 2009). Intact seaweed can be used to demonstrate the partial extracellular location of the vanadium-dependent BPO. When, for example, H₂O₂ and bromide are added to Corallina fronds, A. nodosum (Wever et al., 1991) or L. digitata (Borchardt et al., 2001) immersed in seawater, bromination of phenol red is observed. It is also possible to show that the K _m for bromide is larger than the concentration of bromide in seawater (about 1   mM) by increasing the concentration of bromide to 100   mM. This gives some insight in enzyme kinetics. Further with simple spectrophotometers the initial rates of bromination of the dye monitored at 590   nm can be used to determine an apparent pH optimum for the VBPO. The phenol red assay has also been used in high-throughput screening of mutant libraries of VCPO (Hasan et al., 2006).

2.4.2.2 Monochlorodimedone Assay

To quantitatively determine the halogenating activity of these enzymes the bromination or chlorination of monochlorodimedone is frequently used (De Boer & Wever, 1988). Monochlorodimedone is a 1,3-diketone (Fig. 4) that contains an activated carbon atom that is brominated or chlorinated by HOX or X₂ to form the dihalogenated compound.

Fig. 4. The monochlorodimedone assay is widely used to detect quantitatively halogenating activity. Monochlorodimedone is a 1,3-diketone with an activated carbon atom that is brominated or chlorinated by HOBr or HOCl, respectively, to form the dihalogenated compound. The loss of absorbance of monochlorodimedone is recorded at 290   nm.

By monitoring the loss of absorbance of monochlorodimedone spectrophotometrically at 290   nm (ɛ  =   20   mM ^{−   1}  cm^{−   1}) the brominating or chlorinating activity can be assessed. At neutral or slightly alkaline pH values the H₂O₂ consumption is quantitative with respect to MCD formation. In most cases the assay is carried out in 100   mM citrate buffer (pH 4.5–6.6), 100   mM KBr, 2   mM H₂O₂, and 50   μM monochlorodimedone. This yields a solution with an absorbance of 1 at 290   nm. The exact conditions depend on the type of VHPO. For a new (putative) VHPO a wide range of halide concentrations and multiple halides need to be tested. Also after mutation of a wild-type enzyme kinetics parameters may shift dramatically. In some cases, testing with and without added vanadate (100   μM Na₃VO₄) in the assay may reveal important aspects like strength of cofactor binding. At a pH value of 2.75 the extinction coefficient of MCD drops to 12.2   mM ^{−   1}  cm^{−   1} (Hager, Morris, Brown, & Eberwein, 1966). It is important to correct for this to prevent underestimation of activity at low pH; in many articles, it is unclear whether data have been corrected for this.

Note that at higher pH values singlet oxygen is formed due to a competing reaction of H₂O₂ with hypohalous acid (Eq. 4).

(4) $\mathrm{HOX}+{\mathrm{H}}_2{\mathrm{O}}_2\to {}^1{\mathrm{O}}_2+{\mathrm{H}}_2\mathrm{O}+\mathrm{HX}$

This results in a decrease in the rate of bromination or chlorination of monochlorodimedone (Soedjak, Walker, & Butler, 1995). This is in particular the case when the activity measurements are carried out at higher pH values and or high H₂O₂ levels. In addition, there are some pitfalls with the MCD assay. The observed decrease in absorbance at 290   nm may give false possible results (Wagner, Molitor, & Konig, 2008), and another assay should be used to confirm halogenating activity. It is possible to check this, for example, by using the phenol red assay or use of dimedone instead of monochlorodimedone. Bromination of dimedone will result in an absorbance increase due to formation of monobromodimedone which is subsequently converted into dibromodimedone which has no absorbance at 290   nm (Wever, R., unpublished). A disadvantage of the MCD assay is that a spectrophotometer has to be used with a spectral range in the near UV. Impurities in the seaweed extracts absorbing in the UV may interfere. Either expensive quartz cuvettes are needed to monitor the absorbance changes or specially designed plastic disposable UV cuvettes. Plastic 96-well plates with thin bottoms specially designed for UV may be the most economic solution. When these are used for high-throughput screening with the MCD assay, the enzyme preparations need to have a certain degree of purity to avoid additional UV absorption. It is not advised to use the monochlorodimedone assay to measure IPO activity since the reactivity of oxidized iodine species toward monochlorodimedone is poor. In addition, the iodinated product is unstable and also ${{\mathrm{I}}_3}^{-}$ formed, which absorbs strongly in the UV region (Leblanc et al., 2006 ). Alternatively, the dye thymol blue may be used to monitor the formation of HOBr. Using this dye ( Verhaeghe, Buisson, et al., 2008) obtained kinetic parameters that are essentially the same as those reported (De Boer & Wever, 1988) using monochlorodimedone. As already pointed out the pH range in which this dye can be used is limited to pH values above 7.0.

2.4.2.3 Notes

1.

Some authors use cyanide as inhibitor in this assay to assess whether the BPO contains heme or vanadate as a prosthetic group. However, it should be noted that HOBr will react fast and effectively with cyanide to yield CNBr (Gerritsen et al., 1993; Heeb et al., 2014). As a result, the bromination of monochlorodimedone is inhibited temporarily until all cyanide has reacted.

2.

This assay will not detect haloperoxidases that halogenate and bind specific substrates, e.g., those bacterial enzymes that are involved in bacterial natural product biosynthesis (Bernhardt et al., 2011). Only those enzymes that liberate HOBr or HOCl in solution will be detected.

3.

Some of the VHPOs are inhibited by excess halide, and as already stated for putative VHPOs a range of halide concentrations should be tested in the assay. For the fungal VCPO from C. inaequalis this inhibition is already quite strong at low halide concentrations.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/S0076687918300934

Klebsiella

YingWu Shi , ... Chun Li , in Beneficial Microbes in Agro-Ecology, 2020

2.1 Underlying principles

Klebsiella spp. grow readily on ordinary media commonly used to isolate Enterobacteriaceae, e.g., nutrient agar, tryptic casein soy agar, bromocresol purple lactose agar, blood agar, as well as more differential plating media for Enterobacteriaceae, such as Drigalski agar, MacConkey agar, eosin-methylene blue agar, and bromo-thymol blue agar. K. pneumoniae and K. oxytoca colonies are lactose positive, more or less dome-shaped, 3–4   mm in diameter after overnight incubation at 30 or 37°C, with a mucoid aspect and sometimes stickiness, depending on the strain and the composition of the medium. K. planticola and K. terrigena colonies are also lactose positive, 1.5–2.5   mm in diameter, dome-shaped, with a weakly mucoid aspect. Enterobacter aerogenes (K. mobilis) colonies often have the same morphology. K. ozaenae, K. rhinoscleromatis, and occasionally K. pneumoniae K1 grow more slowly on the same media, yielding voluminous, rounded, very mucoid, translucent, and confluent colonies in 48   h at 30 or 37°C (Orskov, 1981; Richard, 1982). Similar colonies indistinguishable from those of Klebsiella may be formed by other genera of the Enterobacteriaceae, particularly E. coli mucoid varieties with capsular K. antigens (Kauffmann, 1949; Orskov and Fife-Asbury, 1977).

Almost all Klebsiella strains grow in minimal medium with ammonium ions or nitrate as sole nitrogen source and a carbon source (see following) without growth factor requirement. Some K. pneumoniae K1 isolates require arginine or adenine or both as growth factors. K. pneumoniae subsp. rhinoscleromatis requires arginine and uracil. Growth factor requirements of K. pneumoniae subsp. ozaenae are not fully determined. In these requirements, ornithine can replace arginine.

Klebsiella strains can be conserved at room temperature in meat extract semisolid agar, or at −80°C in a broth medium with 10%–50% (v/v) glycerol, or freeze-dried. K. granulomatis has not been grown axenically in artificial media. In vivo cultures in the yolk sac or brain of chick embryo have been achieved. In vitro cultures use fresh mononuclear cells (Kharsany et al., 1997) or the Hep-2 human epithelial cell line (Carter et al., 1997).

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780128234143000137

Gluconobacter

Mitesh Dwivedi , in Beneficial Microbes in Agro-Ecology, 2020

5 Gluconacetobacter diazotrophicus

Gluconacetobacter diazotrophicus (Yamada et al., 1997) was discovered within sugarcane plants in Alagoas, Brazil, by Cavalcante and Dobereiner (1988). Since then, G. diazotrophicus has been found in places such as Mexico and India and in crops ranging from coffee to pineapple. Because they create phytohormones, Gluconacetobacter diazotrophicus has proven to stimulate other plant's growth. The bacterium was initially named as Saccharobacter nitrocaptans and was later classified under AAB and named Acetobacter diazotrophicus, before being reclassified as Gluconacetobacter diazotrophicus based on 16S ribosomal RNA analysis (Cavalcante and Dobereiner, 1988; Gillis et al., 1989; Yamada et al., 1997). Gluconacetobacter diazotrophicus is a Gram-negative, nonspore forming, nonnodule producing, endophytic nitrogen fixing bacterium. The bacterium is an obligate aerobe with cells measuring 0.7–0.9   μm by 2   μm and appears as single, paired, or chainlike structures when viewed under a microscope. The bacterium's cells have one to three lateral or peritrichous flagella used for motility. G. diazotrophicus is an acid-tolerant bacterium, being capable of growing at pH levels below 3.0; however, its optimum pH for growth is 5.5 (Cavalcante and Dobereiner, 1988; Gillis et al., 1989).

These bacteria produce dark brown colonies on potato agar supplemented with 10% sucrose, and dark orange colonies on a nitrogen-poor medium containing bromo thymol blue ( Cavalcante and Dobereiner, 1988). They require a large amount of sucrose for adequate growth (Dong Zhongmin et al., 1994). These bacteria are capable of growth in sucrose levels up to 30%, while optimum growth is achieved at sucrose levels of 10%. They can grow on high concentration of sucrose due to enzyme levansucrase (Martinez-Fleites et al., 2005) that hydrolyze sucrose into fructose and glucose. In addition to sucrose, G. diazotrophicus is capable of abundant growth on other carbon substrates including d-galactose, d-arabinose, d-fructose, and d-mannose (Cavalcante and Dobereiner, 1988). The growth of G. diazotrophicus under laboratory conditions is primarily achieved through plating on LGIP medium due to the fact that it contains high sugar levels which are very similar to those found within sugarcane (quantities per liter: K₂HPO₄, 0.2   g; KH₂PO₄, 0.6   g; MgSO₄·7H₂O, 0.2   g; CaCl₂·2H₂O, 0.02   g; Na₂MoO₄·2H₂O, 0.002   g; FeCl₃·6H₂O,0.01   g; bromothymol blue in 0.2M KOH, 0.025   g; sucrose, 100   g; yeast extract, 0.025   g; agar, 15   g; 1% acetic acid, pH 5.5) (Cavalcante and Dobereiner, 1988). Their growth on LGIP plates can be visualized as smooth colonies with regular edges. These colonies initially appear semitransparent but become dark orange in color due to their uptake of bromothymol blue from within the medium following complete incubation (Cavalcante and Dobereiner, 1988; Gillis et al., 1989). Faster and more robust growth of G. diazotrophicus can be achieved through the addition of a nitrogen source to the LGIP medium, such as 10   mM NH₄(SO₄)₂. Other media capable of sustaining G. diazotrophicus growth include but are not limited to DYGS, C2, ATGUS, modified potato, SYP, AcD, GYC, and EYC media (da Silva-Froufe et al., 2009). The genome that was found to be closely sequenced with Gluconacetobacter diazotrophicus was the Pal5 genome (Eskin et al., 2014). The PAL5 genome yielded one circular chromosome (3,944,163 bp) with a G-C content of 66.19% and two plasmids pGD01 (38,818 bp) and pGD02 (16,610 bp). Overall, the genome contains 3864 putative coding sequences (CDS) (Bertalan et al., 2009).

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780128234143000253

AEROMONAS | Detection by Cultural and Modern Techniques

B. Austin , in Encyclopedia of Food Microbiology (Second Edition), 2014

Commonly Used Media

•

Aeromonas (Ryan's) agar: 0.2% (w/v) l-arginine hydrochloride, 0.3% (w/v) bile salts no. 3, 0.08% (w/v) ferric ammonium citrate, 0.25% (w/v) inositol, 0.15% (w/v) lactose, 0.35% (w/v) l -lysine hydrochloride, 0.5% (w/v) proteose peptone, 0.5% (w/v) sodium chloride, 1.067% (w/v) sodium thiosulfate, 0.3% (w/v) sorbose, 0.375% (w/v) xylose, 0.3% (w/v) yeast extract, 1.25% (w/v) agar, 0.004% (w/v) bromthymol blue, 0.004% (w/v) thymol blue, 5  mg   l⁻¹ ampicillin; pH 8.0; dissolve by boiling; autoclaving is not required. Aeromonas forms dark-green colonies of 0.5–1.5   mm in diameter with dark centers.

•

Alkaline peptone water (APW): 1% (w/v) peptone, 1% (w/v) sodium chloride; pH 8.5–9 (typically at pH 8.5).

•

Ampicillin–dextrin agar (ADA): 1% (w/v) dextrin, 0.01% (w/v) ferric chloride hexahydrate, 0.02% (w/v) magnesium sulfate heptahydrate, 0.2% (w/v) potassium chloride, 0.3% (w/v) sodium chloride, 0.5% (w/v) tryptose, 0.2% (w/v) yeast extract, 1.5% (w/v) agar, 0.004% (w/v) bromthymol blue, 10   mg   l⁻¹ ampicillin, 100   mg   l⁻¹ sodium deoxycholate; pH 8.0. Aeromonas spp. develop as yellow, circular, convex colonies.

•

Bile salts–brilliant green agar (BBG): 1% (w/v) proteose peptone, 0.5% (w/v) Lab Lemco beef extract, 0.5% (w/v) sodium chloride, 0.85% (w/v) bile salts no. 3, 1.5% (w/v) agar, 0.000033% (w/v) brilliant green, 0.0025% (w/v) neutral red; pH 7.2; dissolve by heating; autoclaving is not required. Aeromonas produces whitish colonies on this medium.

•

Bile salts–brilliant green–starch agar (BBGS): 1% (w/v) proteose peptone, 0.5% (w/v) Lab Lemco beef extract, 0.5% (w/v) sodium chloride, 0.5% (w/v) bile salts, 1% (w/v) soluble starch, 1.5% (w/v) agar, 0.005% brilliant green; pH 7.2; dissolve by heating; autoclaving is not required. After flooding with Lugol's iodine, putative Aeromonas may be visualized by the presence of clearing (indicative of starch degradation) around the colonies.

•

Meso-inositol–xylose agar (MIX): 0.01% (w/v) ammonium ferric citrate, 0.2% (w/v) potassium chloride, 0.3% (w/v) sodium chloride, 0.02% (w/v) magnesium sulfate heptahydrate, 1% (w/v) meso-inositol, 0.3% (w/v) yeast extract, 0.15% (w/v) bile salts no. 3, 0.5% (w/v) xylose, 1.5% (w/v) agar, 0.0005% (w/v) bromthymol blue, 20   mg   l⁻¹ ampicillin; pH 7.2. Aeromonas produces convex, circular blue–green colonies.

○

Modified bile salts irgasan brilliant green agar (mBIBG): 0.5% (w/v) meat extract, 0.5% (w/v) proteose peptone, 1% (w/v) soluble starch, 0.58% (w/v) bile salts no. 3, 0.544% (w/v) sodium thiosulfate, 0.0005% (w/v) irgasan, 0.0005% (w/v) brilliant green, 0.0025% (w/v) neutral red, 1.15% (w/v) agar; pH 8.7. Aeromonas develop as purple colonies after incubation at 37   °C for 24   h.

•

Peptone–beef extract–glycogen agar (PBG): 1% (w/v) beef extract, 0.5% (w/v) glucose, 1% (w/v) peptone, 0.5% (w/v) sodium chloride, 0.004% (w/v) bromthymol blue, 1.5% (w/v) agar, and 2% (w/v) agar for overlay. Presumptive Aeromonas appear as yellow colonies with yellow haloes in the otherwise green medium. Ellipsoidal colonies may be seen if they are buried in the medium.

•

Pril ampicillin dextrin ethanol agar (PADE): 1% (w/v) tryptose, 0.2% (w/v) yeast extract, 1.5% (w/v) dextrin, 0.02% (w/v) Pril, 0.02% (w/v) MgSO₄·7H₂O, 0.01% (w/v) FeCl₃·6H₂O, 0.005% (w/v) bromothymol blue, 0.005% (w/v) 0.005% (w/v) thymol blue, 1.5% (w/v) agar, autoclave at 110   °C for 20   min before adding 10   ml ampicillin (3   mg   ml⁻¹), 10   ml sodium deoxycholate (10   mgml⁻¹), 1% (v/v) ethanol; pH 8.6. Aeromonas develop as yellow colonies after incubation at 37   °C for 24   h.

•

Rimler Shotts medium (RS): 0.05% (w/v) l-lysine hydrochloride, 0.65% (w/v) l-ornithine hydrochloride, 0.35% (w/v) maltose, 0.68% (w/v) sodium thiosulfate, 0.03% (w/v) l-cysteine hydrochloride, 0.003% (w/v) bromthymol blue, 0.08% (w/v) ferric ammonium citrate, 0.1% (w/v) sodium deoxycholate, 0.0005% (w/v) novobiocin, 0.3% (w/v) yeast extract, 0.5% (w/v) sodium chloride, 1.35% (w/v) agar; pH 7.0: After boiling to dissolve the ingredients, autoclaving is not required. Aeromonas develop as yellow colonies after incubation of spread plates of RS at 30   °C for 24   h.

•

Rippey–Cabelli agar (mA): 0.1% (w/v) ferric chloride hexahydrate, 0.02% (w/v) magnesium sulfate heptahydrate, 0.2% (w/v) potassium chloride, 0.3% (w/v) sodium chloride, 0.5% (w/v) trehalose, 0.5% (w/v) tryptose, 0.2% (w/v) yeast extract, 1.5% (w/v) agar, 0.004%(w/v) bromthymol blue, 1% (v/v) ethanol, 20   mg   l⁻¹ ampicillin, 100   mg   l⁻¹ sodium deoxycholate, pH 8.0. Aeromonas spp. develop as yellow, circular, convex colonies.

•

Starch–ampicillin agar (SAA): 0.1% (w/v) beef extract, 1% (w/v) proteose peptone no. 3, 0.5% (w/v) sodium chloride, 0.1% (w/v) starch, 1.5% (w/v) agar, 25   mg   l⁻¹ of phenol red, 10   mg   l⁻¹ of ampicillin. Putative Aeromonas colonies are 3–5   mm in diameter, and are yellow to honey pigmented. After flooding the plates with full or half strength Lugol's iodine, Aeromonas colonies will be surrounded by a clear zone, indicating such hydrolysis.

•

Tryptone–soya–ampicillin broth (TSAB): tryptone soya broth containing 30   mg   l⁻¹ ampicillin.

•

Xylose–deoxycholate–citrate agar (XDCA): 1.25% nutrient broth no. 2, 0.5% (w/v) sodium citrate, 0.5% (w/v) sodium thiosulfate, 0.1% (w/v) ferric ammonium citrate (brown), 0.25% (w/v) sodium deoxycholate, 1.2% (w/v) agar, 1% (w/v) xylose, 0.0025% (w/v) neutral red; pH 7.0; dissolve by heating; autoclaving is not required. Aeromonas develop as colorless colonies.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780123847300000057

Rabeprazole: A comprehensive profile

Ahmed H. Bakheit , ... Salem Albraiki , in Profiles of Drug Substances, Excipients and Related Methodology, 2021

5.2.2.1 Spectrophotometry

El-Gindy et al. [28] proposed three methods for evaluating Rabeprazole in the presence of its degradation products. One of these methods was based on the first derivative of the spectra ratio (¹DD), which relied on the measurement of amplitudes at 310.2   nm. The pH-rate profile of rabeprazole degradation in Britton–Robinson buffer solutions within pH range 3–11 was further investigated. In Britton–Robinson buffer solution pH 7, however, the activation energy of rabeprazole degradation was determined. The calibration range of rabeprazole with the ratio first derivative of the spectra ratio was 10–30   μg/mL, the detection limit was 0.019, and the quantitation limit was 0.058.

Patel et al. [29] developed a spectrophotometric method for determination of rabeprazole in pharmaceutical bulk dosage form. The method depended on the ion-pair complexes formation of rabeprazole with four dyes, viz. bromo thymol blue, bromocresol green, bromophenol blue and bromocresol purple in solution of acidic buffer, afterwards their chloroform extraction. The absorption of the organic layer was measured against the corresponding blank reagent at the respective wavelength of the maximum absorbance. The method was evaluated statistically and found to be accurate and precise. Phosphate buffer (pH 2) and bromocresol green dye phosphate buffer gave the maximum absorption of rabeprazole at 454  nm. The Beer law was found within the concentration range of 10–100   μg/mL.

A spectrophotometric derivative method for evaluating Rabeprazole sodium was developed and validated by Garcia et al. [30] in drug formulations. The procedure was performed with water as a diluent (pH 10.0). The derivative spectra of the first order were obtained at N  =   5, Δλ   =   4.0   nm, and determinations were made at wavelength 304   nm. In the present of drug excipients, the procedure showed a good linearity of 6.0 and 18.0   μg/mL^{−   1}, and the accuracy was 99.15%.

Sabnis et al. [31] developed a spectrophotometric ratio method of spectra derivatives for simultaneous determination of rabeprazole sodium and itopride hydrochlorides in theirs dosage forms. Amplifications were chosen for the first derivatives of the respective at 231   nm (minimum) and 260   nm ratio range. Linearity, accuracy and precision, were validated. The Beer law was found to be within the 4–20   μg/mL concentration range for rabeprazole.

Gul et al. [32] reported a UV spectrophotometric method for determination of rabeprazole in bulk using methanol as solvent. The solution was scanned at 200–400   nm with a maximum absorption at 284   nm. The method was validated according to ICH guidelines. Linearity was in the 12–18   μg/mL range. A recovery was 99.86–100.14%. The %RSD for repeatability was 0.628% and for precision and inter was found at 0.488–0.77%. This method indicated that rabeprazole was interactive with clorazepate dipotassium at pH 7.4.

Garcia et al. [33] developed a UV spectrometric ultraviolet (UV) method for determination of rabeprazole in pharmaceutical formulation and compared the method with a previously accepted capillary electrophoresis (CE) method. The proposed method was applied using water (pH 10.0) as diluted at 291   nm. The method showed a good linearity (r  =   0.9997) in the concentration range of 6.0–18.0   μg   mL^{−   1}.

Gowri et al. [34] developed two new methods for estimation of Rabeprazole in bulk and tablet dosage forms. In the first method, a Folin-Ciocalteu reagent and sodium hydroxide were used. A Folin-Ciocalteu reagent was reduced by Rabeprazole, in the presence of sodium hydroxide, and blue colored chromogen was formed at a maximum absorption of 640   nm. The linearity range was obtained at concentration of 1–8   μg/mL and a correlation coefficient (r²) was found to be 0.9983. In the second method, the ferric nitrate was reduced by rabeprazole. The product ferrous ions reacted with 2,2,bipyridine, showing maximum at 522   nm. The linearity was obtained at range of concertation of 2–16   μg/mL with a correlation coefficient (r²) of 0.9994. These two methods were validated according to the ICH guidelines and they can be applied to determination of rabeprazole in bulk and dosage form.

A simultaneous UV spectrophotometric method for determination of Famotidine and Rabeprazole sodium in the laboratory mixture was developed and validated by Dsouza et al. [35]. The proposed method was performed at a maximum absorption of 263   nm for Famotidine and 284   nm for Rabeprazole sodium. The method obeyed Beer's law in the concentration range of 5–30   μg/mL for both famotidine and rabeprazole.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/S1871512520300157

Nanomaterials-based optoelectronic noses for food monitoring and classification

Jose Vicente Ros-Lis , ... Ramón Martínez-Máñez , in Nanobiosensors, 2017

7 Determination of Blue Cheese Origin

Five cheeses (four blue cheeses and Cheddar) were used in this study. Three of the selected blue cheeses were from diverse origin (Blue Stilton, Rochefort, and blue cheese with leaves) and the fourth was blue cheese spread. Cheddar was tested as control (a nonblue cheese). It has been reported that times of up to 16 h or temperatures above room temperature are necessary to reach the equilibrium in the headspace (Cakmakci et al., 2013; Frank et al., 2004). So although the array may be suitable as a fast screening method to classify or identify blue cheeses, the array response may vary with time, and this factor should be controlled to avoid misclassification. Thus in this study time was controlled, and color data from the array was taken at 0.5, 1.5, 3.5, 5.5, and 8.5 h after packaging. The optoelectronic array for the analysis of the origin of blue cheeses was composed by five sensing materials (Table 1.1 ). They consist of three pH indicators (br-cresol purple, m-cresolsulfonphthalein, thymol blue), which were combined with two support materials: aluminium oxide (basic) and MCM-41 (slightly acid). The objective was to design an optoelectronic nose that was as simple as possible that could be used by consumers and retailers. For such purposes pH indicators offer advantages over other indicators since they are cheap and commercially available, and can be easily incorporated to support ensuring array scalability.

Color differences were analyzed by PCA. Lab-based color data generated by the responses of the five sensing materials used in the chromogenic array was used to perform the analysis and an autoscaled technique was applied to the data set. PCA study at 30 min revealed a high degree of dispersion among the independent dimensions. PC1 contained only 37.03% of data variance and the first eight PCs were required to account for 95% of variance. The first three components, representing the main part of total variance (70.60%), were plotted on the x, y, and z axes as a method to project data to a three-dimensional hyperplane. Fig. 1.7 shows the resulting PCA for the five cheeses (three replicates) 30 min after packaging. As observed, all cheeses clustered in a completely different zone, confirming that the chromogenic array generates a characteristic fingerprint for each blue cheese.

Figure 1.7. The principal components analysis (PCA) score plot of the diverse cheeses using the data obtained 0.5 h after packaging.

Data are shown for three different trials: (♦) Blue Stilton; (▪) blue cheese with leaves; (▴) Cheddar; (▾) Roquefort; (⋆) blue cheese spread.

Reproduced from Zaragozá et al. (2015), with permission of Elsevier.

Moreover the data set was also analyzed by PLS-DA. The output of PLS-DA consists in assigning each sample to one of the predefined categories that, in line with our general purpose of obtaining an array capable of making classifications between blue cheeses, was each one of the cheeses. Five classes (one for each blue cheese and one for the Cheddar control) were fixed and the model was applied to the data collected after only 30 min (see first entry on Table 1.6). An average of 67% of correct prediction was obtained at 30 min. Although 67% of proper assignment is in line with the results obtained with other technologies for the same purpose (Kulmyrzaev et al., 2008), it is probably too low for practical use. Classifications close to 100% are required to ensure users' confidence in this technology.

Table 1.6. Percentage of Samples Correctly Classified by the PLS-DA Model Using Five Classes (Cheddar, Roquefort, Blue Stilton, Blue Cheese With Leaves, and Blue Cheese Spread): Time Indicates That Color Data up to the Time Spent in the PLS-DA Model

Time	LV	Blue Stilton	Blue Cheese With Leaves	Roquefort	Blue Cheese Spread	Cheddar
0.5 h	4	0%	67%	67%	100%	100%
1.5 h	4	33%	33%	100%	100%	100%
3.5 h	4	33%	67%	100%	67%	100%
5.5 h	4	100%	100%	100%	100%	100%
8.5 h	4	100%	100%	100%	100%	100%

We must take into account that we are using only five sensing materials to classify five cheeses. Cross-sensitive sensors tend to generate a limited amount of information, which is the reason why, in general, the arrays tend to require a large number of indicators. Since we were interested in keeping the array as simple as possible, and as a strategy to increase the information and the number of independent variables, the PLS-DA model of our chromogenic array was fed with not only the data collected at a certain time, but also with the data measured up to the labelled time (ie, for 1.5 h, the data measured at 0.5 and 1.5 were incorporated into the model). As can be seen on Table 1.6, as more information was incorporated, the prediction ability improved from 67% at 30 min to 100% at 5.5 h and 8.5 h. This perfect prediction at 5.5 h confirms the capacity of the chromogenic array to classify blue cheeses and opens up the use of optoelectronic noses as a real, easy-to-use, and rapid classification tool for cheeses.

Given our interest in developing a tool as quickly as possible and the above-reported results at 30 min, also a PLS-DA model with a lower number of classes was run. If three classes are used: (1) Roquefort, (2) Cheddar, and (3) the remaining blue cheeses. All three classes were correctly classified by 30 min (100% of correct assignments), which suggests that the array can be a useful rapid Roquefort classification test.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B9780128043011000011

LIQUID CHROMATOGRAPHY | Affinity Chromatography

D.S. Hage , in Encyclopedia of Analytical Science (Second Edition), 2005

Applications of Affinity Chromatography

Preparative Applications

The earliest use of affinity chromatography and its most popular application is in the purification of proteins and other biological agents. The use of this method in enzyme purification is particularly important, with hundreds to thousands of applications having been reported in this field alone. Ligands used for this purpose include enzyme inhibitors, coenzymes, substrates, and cofactors. For instance, nucleotide mono-, di-, and triphosphates can be used for the purification of various kinases, NAD has been used to isolate dehydrogenases, and RNA or DNA has been used for the preparation of polymerases and nucleases.

Antibodies have also been popular ligands for the purification of biological compounds. There are now thousands of examples of immunoaffinity methods that have been developed for the isolation of hormones, peptides, enzymes, recombinant proteins, receptors, viruses, and subcellular components. In addition, immobilized antigens are frequently used to isolate specific types of antibodies. A more general purification scheme for antibodies can be obtained by using antibody-binding proteins like protein A and protein G. These latter ligands have the ability to bind to the constant region of many types of immunoglobulins. Both protein A and protein G have their strongest binding to immunoglobulins at or near neutral pH but readily dissociate from these when placed into a lower pH buffer.

Dye–ligand affinity chromatography is often used in large-scale protein and enzyme purification, with over 500 such compounds having been isolated by this technique. In this method, an immobilized synthetic dye is used that binds to the active site of a target by mimicking the structure of its substrate or co-factor. The most common dye used for this purpose is Cibacron Blue 3G-A (see Figure 2 ). Other dyes used include Procion Blue MX-3G or MX-R, Procion Red HE-3B, Thymol Blue, and Phenol Red. Although these ligands were originally discovered on a trial and error basis, recent work in the area of biomimetic affinity chromatography has used computer modeling and three-dimensional protein structures to develop dyes that compliment the binding pockets of specific target proteins.

Figure 2. Structure of Cibacron Blue 3G-A, a stationary phase often used in dye–ligand affinity chromatography.

Analytical Applications

Although affinity chromatography was originally created as a preparative method, the past few decades have seen this method also become an important tool in analytical applications. Table 4 summarizes some strategies that can be employed in these analyses and gives examples of representative applications. The simplest format for using affinity chromatography in analysis involves the traditional step gradient mode, as shown in Figure 1. The advantages of using this approach in analytical applications, particularly when performed by HPLC, include its speed, relative simplicity, and good precision.

Table 4. Analytical applications of affinity chromatography ^a

General application	Examples of analytes
Direct detection	Anti-idiotypic antibodies, antithrombin III, bovine growth hormone, fibrinogen, fungal carbohydrate antigens, glucose tetrasaccharide, glutamine synthetase, granulocyte colony stimulating factor, group A-active oligosaccharides, human serum albumin, immunoglobulin G, immunoglobulin E, interferon, interleukin-2, lymphocyte receptors, β 2-microglobulin, tissue-type plasminogen activator, transferrin
Offline affinity extraction	Aflatoxin, albuterol, benzodiazepines, cytokinins, fumonisin, human chorionic gonadotropin, ivermectin and avermectin, nortestosterone, ochratoxin A, oxytocin, phenylurea herbicides, sendai virus protein, trenbolone, triazine herbicides
Online affinity extraction	Aflatoxin M1, β-agonists, α 1-antitrypsin, atrazine, atrazine metabolites, benzylpenicilloyl-peptides, bovine serum albumin, carbendazim, carbofuran, chloramphenicol, clenbuterol, cortisol, dexamethasone, diethylstilbestrol, digoxin, estrogens, hemoglobin, human epidermal growth factor, human growth hormone variants interferon α-2, LSD, lysozyme variants
Chromatographic immunoassays	Adrenocorticotropic hormone, α-amylase, atrazine/triazines, 2,4-dinitrophenyl lysine, human chorionic gonadotropin, human serum albumin, immunoglobulin G, isoproturon, parathyroid hormone, testosterone, theophylline, thyroid stimulating hormone, thyroxine, transferrin, transferrin, trinitrotoluene

a

The information in this table is based on data provided in Hage (1998).

Affinity extraction is another approach that can be used for solute detection. In this method, an affinity column is used for the removal of a specific solute or group of solutes from a sample prior to their determination by a second method. This employs the same operating scheme as shown in Figure 1, but now involves combining the affinity column either offline or online with some other method for the actual quantitation of analytes. This often involves the use of antibodies as ligands, but other binding agents can also be employed.

Offline extraction is the easiest and most common way for combining affinity columns with other analytical techniques. This typically involves the use of antibodies that are immobilized and packed into a disposable syringe or solid-phase extraction cartridge. After conditioning the affinity column with the necessary application buffer or conditioning solvents, the sample is applied and undesired sample components are washed away. An elution buffer is then applied and the retained target is collected. If desired, the collected fraction can be analyzed directly or first dried down and reconstituted in a solvent that is more compatible with the method to be used for quantitation.

Online affinity extraction can also be used. A typical scheme for performing online immunoextraction with reversed-phase liquid chromatography (RPLC) is shown in Figure 3. This involves injecting the sample onto the immunoextraction column, with this column later being switched online with a RPLC column. An elution buffer is then applied to dissociate any retained analyte, which will be captured and reconcentrated at the head of the RPLC column. After all solutes have left the immunoaffinity column, this column is switched back offline and regenerated by passing through the initial application buffer. Meanwhile, the RPLC column is developed with either an isocratic or gradient elution scheme that uses a mobile phase with increased organic modifier content. As the solutes elute through the RPLC column, they are monitored and quantitated through the use of an online detector.

Figure 3. A general scheme for interfacing an immunoextraction column with reversed-phase liquid chromatography (RPLC). The two circles represent six-port switching valves, with the solid and dashed lines showing the flow of sample and solvents in each of the two positions. The operation of this system is described in the text.

A third way in which affinity chromatography has been used in analytical applications has been in the area of chromatographic, or flow-injection, immunoassays. This method is used in determining trace analytes that do not directly produce a readily detectable signal. The competitive binding assay is the most common format used in performing such an assay. This is generally accomplished by mixing the sample with a fixed amount of a labeled analyte analog (i.e., the 'label') and simultaneously or sequentially injecting these onto an immunoaffinity column that contains a limiting amount of antibody. Immunometric assays have also been performed on chromatographic systems. For instance, in a sandwich immunoassay, two different types of antibodies are used (see Figure 4). The first of these two antibodies is attached to a solid-phase support and used to extract the analyte from samples. The second antibody contains an easily measured tag and serves to place a label on the analyte, thus allowing it to be quantitated.

Figure 4. Detection of a trace hormone (parathyrin) in human plasma by a chromatographic sandwich immunoassay. The results in (A) show the injection spikes and nonretained fractions for four sequential injections of human plasma samples with increasing parathyrin levels. The results in (B) show the response due to the retained parathyrin. This plot was obtained from Hage and Kao (1991).

Biophysical Applications

Besides its use in separating and quantitating sample components, affinity chromatography can also be employed as a tool for studying solute–ligand interactions. This approach is called analytical, or quantitative, affinity chromatography. Using this technique, information can be obtained regarding the equilibrium and rate constants for biological interactions, as well as the number and types of sites involved in these interactions.

Information on the equilibrium constants for a solute–ligand system can be acquired by using the methods of zonal elution or frontal analysis. Zonal elution involves the injection of a small amount of solute onto an affinity column in the presence of a mobile phase that contains a known concentration of competing agent. The equilibrium constants for the ligand with the solute (and competing agent) can then be obtained by examining how the solute's retention changes as the competing agent's concentration is varied (see Figure 5). This method has been used to examine a number of biological systems, such as enzyme-inhibitor binding, protein–protein interactions, and drug–protein binding. Frontal analysis is used in a similar manner but is performed by continuously applying a known concentration of solute to the affinity column. The moles of analyte required to reach the mean point of the resulting breakthrough curve is then measured and used to determine the equilibrium constant for solute–ligand binding.

Figure 5. An example of a zonal elution experiment, in which small injections of l-tryptophan are made on to an immobilized human serum albumin column in the presence of increasing amounts of phenytoin in the mobile phase.

Information on the kinetics of solute–ligand interactions can also be obtained using affinity chromatography. A number of methods have been developed for this, including techniques based on band-broadening measurements, the split-peak effect, and peak decay analysis. These methods are generally more difficult to perform than equilibrium constant measurements but represent a powerful means for examining the rates of biological interactions. Systems studied by these techniques have included the binding of lectins with sugars, protein A, or protein G with immunoglobulins, antibodies with antigens, and drugs with serum proteins. The recent creation of commercial sensors for biointeraction studies is one result of such work.

Read full chapter

URL:

https://www.sciencedirect.com/science/article/pii/B0123693977003216

carreonhielturry.blogspot.com

Source: https://www.sciencedirect.com/topics/agricultural-and-biological-sciences/thymol-blue

Share this post