Quantitative assessment of phase composition and morphology of two-phase gelatin–pectin gels using fluorescence microscopy

Food Hydrocolloids 14 (2000) 579–590 www.elsevier.com/locate/foodhyd

Quantitative assessment of phase composition and morphology of twophase gelatin–pectin gels using fluorescence microscopy T.S. Nordmark, G.R. Ziegler* Department of Food Science, The Pennsylvania State University, 116 Borland Laboratory, University Park, PA 16802 USA Received 14 January 2000; revised 2 June 2000; accepted 2 June 2000

Abstract A technique for quantitative determination of the concentrations of polysaccharide and protein in two-phase mixtures by fluorometry has been developed and compared with chemical analysis. In the first case, a general method for fluorescent labeling of carbohydrate polymers was developed. For the latter purpose, two micro-assays were developed on the basis of recent polymer macro-assays. A blend of lowmethoxyl pectin and gelatin B was used as a model system. The commercial components were subjected to multi-step purification procedures, and phase separation was initiated by the addition of NaCl to aqueous solutions containing the two polymers. Samples were withdrawn for microscopy after various holding times at 60⬚C. Tie-lines were determined using both the fluorescent and chemical methods. The results from these methods were in fair agreement with each other and with literature data. A three-phase region was discovered in the pseudo-ternary phase diagram. The morphology of double labeled gels was also studied in two and three dimensions using confocal scanning laser microscopy. The results show promise for the quantitative assessment of phases that contain carbohydrate polymers and in the study of morphological changes that occur during thermo-mechanical processing. 䉷 2000 Elsevier Science Ltd. All rights reserved. Keywords: Gum; Low-methoxyl pectin; Gelatin; Fluorescence; Morphology; Phase diagram

1. Introduction Structure-forming polysaccharides and proteins provide desired functional properties to a wide range of foods (Kinsella, Rector, & Phillips, 1994). The characteristics of blends of such hydrocolloids both in the liquid and gel states have lately been the subject of an increasing number of investigations because of the prospect of discovering useful synergistic effects (Ipsen, 1995). This research has begun to demonstrate how physical properties of blends can be related to phase morphology and quantitative relationships can be established (Owen & Jones, 1998). A number of different analytical approaches have been employed to elucidate structure–function relationships (BeMiller, 1996; Dickinson & McClements, 1995; Kalab, Allan-Wojtas, & Miller, 1995; Ross-Murphy, 1994). Direct determination of phase-composition is an obvious major target for future research on biopolymer co-gels (Kasapis, Morris, Norton, & Clark, 1993b). The present work was undertaken to expand the use of fluorescence-based analytical methods to research on phaseseparated food materials and the relation of morphological * Corresponding author. Tel.: ⫹1-814-863-2960; fax: ⫹1-814-863-6132. E-mail address: [email protected] (G.R. Ziegler).

and compositional features to rheological properties. Fluorescence and fluorescence microscopy in biology are expounded upon by Ichinose, Schwedt, Schnepel, and Adachi (1991), Ploem (1993), and Rost (1991). Applications of fluorescence in food research have been recently reviewed (Blonk & van Aalst, 1993; Strasburg & Ludescher, 1995; Vodovotz, Vittadini, Coupland, McClements, & Chinachoti, 1996) and the properties of gelatin– pectin gels have been the focus of research (Al-Ruqaie, Kasapis, & Abeysekera, 1997; DeMars, 1995; Gubenkova, Somov, & Shenson, 1988). Gelatin is derived from denatured collagen that has been further processed, and the dominating amino acids are glycine, proline, and hydroxyproline. Thermoplastic gels are formed upon cooling, and a blend of fine and coarse networks can be found (Ziegler & Foegeding, 1990). Gelatins of type B have their isoelectric points close to pH 4.9 and below this may form complex coacervates with negatively charged polysaccharides. Pectic substances are linear, partly methylesterified polygalacturonic acid chains, where neutral sugars like rhamnose may be present as side chains or inserted in the main chains (da Silva & Goncalves, 1994). Both smooth and hairy chains, the latter with side chains of arabinogalactan or other oligosaccharides, may exist (Aman & Westerlund, 1996). The galacturonic acid residues

0268-005X/00/$ - see front matter 䉷 2000 Elsevier Science Ltd. All rights reserved. PII: S0268-005 X( 00)00 037-0

580

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

contain vicinal diols, which we employ in the protocol for fluorescent labeling. There are fewer alternatives for the fluorescent labeling of polysaccharides than for the labeling of proteins, in particular for general and quantitative purposes. Food proteins are frequently labeled with FITC (fluorescein isothiocyanate), Rhodamine, or Texas Red (Blonk & van Aalst, 1993). Traditional dyes such as Calcofluor White and Anilin Blue often attach to specific carbohydrate residues or linkages (Fulcher, Faubion, Ruan, & Miller, 1994). The lectins from pea tree (Caragana arborescens) have been shown to bind agarose beads and can be marked with FITC (EY Laboratories, 1990). Polysaccharide side chains can be made more reactive by using transferases (Brossmer & Gross, 1994; Gahmberg & Tolvanen, 1994) or by using galactose oxidase (EC 1.1.3.9), which predominantly, but not always exclusively, acts upon terminal, non-reducing dgalactose residues (Goudsmit, Matsuura, & Blake, 1984; Mazur, 1991; Wilchek & Bayer, 1987). In this paper we present a protocol for covalent labeling of pectin and many other carbohydrate polymers with the fluorescent probe BODIPY FL hydrazide. This probe has recently been used for the quantification of progesterone and other 3-keto steroids by HPLC (Katayama et al., 1998). We labeled gelatin covalently with the succinimidylester of carboxytetramethylrhodamine by slightly modifying an existing protocol for labeling of globular proteins that contain aliphatic amines. This probe has lately been conjugated with peptides for inclusion and detection in degradable poly(lactic acid) (PLA) microspheres (Brunner, Minamitake, & Gopferich, 1998). The phase behavior of mixed polysaccharides and proteins has been investigated by employing centrifugation of the phases, chemical analyses, osmotic pressure measurements, light scattering, FTIR, and turbidimetry (Antonov, Lashko, Glotova, Malovikova, & Markovich, 1996; Clewlow, Rowe, & Tombs, 1995b; Durrani, Prystupa, Donald, & Clark, 1993; Vinches, Parker, & Reed, 1997). Improvements in most of these techniques cannot compensate for the difficulties that arise due to increased viscosity in more concentrated mixes. Accordingly, there is an interest in the development of methods that could be used in situ, e.g. Blonk, van Eendenburg, Koning, Weisenborn, and Winkel (1995) discussed the use of confocal scanning laser microscopy in combination with fluorescent double-labeling of alginate and caseinate (max. 2 and 10%, respectively) in the liquid state. The objective of this work was the development of a method for the in situ measurement of phase composition and morphology in highly viscous biopolymer mixtures.

remove electrolytes and particles. Commercial low-methoxyl citrus pectin (LM290 NA95 from SKW Biosystems, Inc., Waukesha, WI) with 31.9% degree of esterification and gelatin type B (Sigma Chemical Co., St. Louis, MO) made from bovine skin tissue were used as polymeric raw materials. Polymers were purified from cations and sugars using the following procedure. The pectin and gelatin were dispersed in cold 25 and 40% aqueous ethanol, respectively, diafiltered 4 times, and treated batch-wise with stirred AG50W-X8 (20/50 mesh) cation exchange resin (Bio-Rad Laboratories, Hercules, CA). The desalted pectin and gelatin dispersions were decanted, the resin was washed with aqueous ethanol until clean of polymer, and the used wash liquid was added to the polymer dispersions. The dispersions were then filtered through a 50 mm fritted glass filter funnel, diafiltered with 100 ml aq. ethanol (40 and 25% concentration for gelatin and pectin, respectively), titrated to pH 5.5 (using 100 mM hydrochloric acid or sodium hydroxide), slowly precipitated with ethanol, freeze-dried for 48 h, ground in an analytical mill, and stored in a dessicator at 20⬚C (Berth, 1988; Doner & Douds, 1995; Walter & Sherman, 1983). The fluorescent probes D-2371 (BODIPY FL hydrazide), C-1171 (TAMRA succinimidylester), and T6105 were purchased from Molecular Probes, Inc. (Eugene, OR). Supor-450 polysulfone membrane filters (0.45 mm) from Gelman Sciences (Ann Arbor, MI) were used when filtering the polymer solutions. Warm, gas tight syringes (Hamilton Co., Reno, NV) were utilized for the quantitative transfer of high viscosity polymer solutions at 50⬚C. Disposable 10 DG polyacrylamide size exclusion chromatography (SEC) columns (Bio-Rad Laboratories, Hercules, CA) and Centricon-10 centrifugal concentrators (Amicon, Inc., Beverly, MA) were used in the protocols for fluorescent labeling. An LSM 410 inverted Laser Scan Microscope (Carl Zeiss, Inc., Thornwood, NY) with argon- and helium– neon lasers and photomultiplier tube detectors was employed for confocal fluorescence microscopy. A Nikon Diaphot 300 inverted fluorescence microscope (Nikon Inc., Melville, NY), a 75 W xenon lamp, and a liquid cooled CCD-camera of type CH 250 (Photometrics Ltd., Tucson, AZ) were used for wide-angle fluorescence microscopy. Cytoseal 60 (Stephens Scientific, Riverdale, NJ) was generally employed as sealant for gels mounted for microscopy. The image processing software was IPLab Spectrum HSU2, v. 2.5.7 (Signal Analytics Corporation, Vienna, VA). An IEC Model CL centrifuge (International Equipment Company, Needham Heights, MA) was employed in the reference experiments of the quantitative study.

3. Methods 2. Materials The water was treated in a NANOpure water purification system (Barnstead/Thermolyne, Inc., Dubuque, IO) to

Purified pectin and gelatin were covalently labeled with fluorescent probes as described below. The concentration of the fluorescent solution was estimated using

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590 Disperse 30 mg purified LM pectin in 2 ml cold deionized water while using a stir bar at high speed and continue stirring at moderate speed for 1 hour Boil for 30 seconds Cool to 60 ¡ C and filter 1 ml Cool to 20 ˚C Add 0.5 ml periodate solution while using a stir bar at moderate speed and continue stirring at low speed for 30 minutes in darkness Add 50 µL of 2 M aq. glycerol while stirring and wait for 5 minutes Filter through a 6 kD SEC column and elute with 10 mM PBS buffer pH 7.4 Concentrate to 1.2 ml by centrifugation at 5,000 g Add 50 µL solvent, stir, and slowly add 33 µL of fluorescent solution Stir at low speed for 3 hours in darkness and overnight in a refrigerator Add 1-2 mg sodium cyanoborohydride and stir at low speed for 40 minutes Filter twice through 6 kD SEC columns and elute with deionized water Concentrate and deaerate to 1.1 ml by centrifugation at 5,000 g

581

3.1. Pectin Pectin was fluorescently labeled with the non-ionic dye BODIPY FL (4,4-difluoro-5,7-dimethyl-4-bora-3a,4adiaza-s-indacene-3-propionylhydrazide), which is more photostable than fluorescein and has high extinction coefficient and quantum yield (Haugland, 1996). Its emission spectrum is reasonably distant from the excitation spectrum of the TAMRA dye, which was used for labeling of gelatin. The hydrazide group forms conjugates with ketogroups (Hermanson, 1996). A fluorochrome concentration of 250 mg/ml in the reaction mixture was chosen (in similarity with the work by Katayma et al., 1998).Reagent solutions. Periodate solution: Take 0.5 ml of a 10 mM stock solution of sodium metaperiodate, and add 1.75 ml deionized water and 65 mg PBS (phosphate buffered saline)-powder as buffer salt (Sigma Chemical Co., St. Louis, MO). Fluorescent solution: Dissolve 5 mg fluorescent probe (D-2371) in 500 ml methanol (Omnisolv from EM Industries, Inc., Gibbstown, NJ) by vortexing. Dimethylformamide (DMF) (ACS Reagent from Sigma Chemical Co., St. Louis, MO) may be employed as solvent if the probe solution will be used within one day. This solvent was used in the qualitative study. Fluorescent labeling: See Fig. 1.

Heat to 60 ˚C; evaporate to 1.0 ml; solution may be stored in a refrigerator

Fig. 1. Fluorescent labeling of LM pectin.

spectrophotometry before the start of each labeling procedure. Purified and labeled polymer was characterized using the micro-assays described below. Disperse 40 mg purified gelatin in 2 ml cold deionized water while using a stir bar at moderate speed and continue stirring at low speed for 1 hour Boil for 30 seconds Cool to 60 - 70 ¡ C and filter 1.3 ml Cool to 40 ˚C Slowly add 130 µL of fresh dissolved fluorescent probe while using a stir bar and continue stirring the covered solution at low speed for 1.5 hours Add 130 µL fresh hydroxylamine solution and stir the covered mix at low speed for 1 hour Filter through a 6 kD SEC column and elute with PBS buffer pH 7.4 Filter through a 6 kD SEC column and elute with deionized water Concentrate and deaerate to 1.1 ml by centrifugation at 5,000 g Heat to 60 ˚C and evaporate to 1.0 ml; the covered solution may be stored in a refrigerator

Fig. 2. Fluorescent labeling of gelatin.

3.2. Gelatin Gelatin was fluorescently labeled with mixed isomers of TAMRA SE (5- and 6-carboxytetramethylrhodamine succinimidylester), which are among the most photostable fluorescent dyes available and emit fluorescent light of high intensity. A protein conjugate made from a succinimidyl ester of TAMRA is considered to be more chemically stable than a conjugate made from the isothiocyanate derivatives commonly used (Haugland, 1996). The overlap between its excitation spectrum and the emission spectrum of the BODIPY FL dye is reasonably small. The CSLM helium–neon laser delivers light at 543 nm, which closely matches the 546 nm excitation wavelength of the TAMRA SE. It may form non-fluorescent dimers when attached to proteins and is more prone to degrade in moist environments (liquid or solid state) than the BODIPY probe is (Molecular Probes, Inc. Eugene, OR). The protocol below was followed.Reagent solutions. Fluorescent solution: Make a solution of 10 mg probe C-1171/ml DMF in a conical microcentrifuge tube and vortex it until the probe is dissolved. Hydroxylamine solution: Dissolve 420 mg dry fresh hydroxylamine hydrochloride in 2 ml deionized water and carefully adjust the pH to 8.5 while stirring. Dilute the solution with water so that the final concentration will be 1.5 M. Fluorescent labeling: See Fig. 2. All treatments of the gelatin solution were carried out in a 40⬚C environment in order to avoid gelation.

582

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

Disperse purified LM pectin and purified gelatin separately in deionized water using a stir bar at high and moderate speed, respectively. Continue to stir at moderate speed for 1 hour Boil each dispersion for 30 seconds and filter through a 0.45 µm membrane Cool each dispersion to 40 - 50 ˚C Mix the dispersions at 40 ˚C Concentrate to approximately final polymer concentrations or slightly less by evaporation at 70 ˚C Adjust to pH 5.5 if required Stir and add a warm 20 % (w/w) solution of sodium chloride Boil for 30 second Evaporate at 60 ˚C until final concentrations (including 1 M NaCl) are obtained Pour into a pre-weighed conical PP centrifuge tube and weigh again Keep the capped tube in a 60 ˚C water bath for 3 hours minus the evaporation time in the previous step Spin at 1,100 g and 60 ˚C for 10 minutes in a preheated centrifuge Immediately cool the tubes briefly in ice-water and refrigerate the tubes for 30 minutes Cleave the tube transversely with a hot knife; weigh the gel containing part Cleave this part longitudinally; place each phase in a separate, pre-weighed graduated tube and weigh the covered tubes and the original tube halves Dilute each phase with 60 ˚C water as required for the polymer micro-assays and remove duplicate samples Analyze the remnants for moisture content by oven drying at 100 ˚C for 2 hours

Fig. 3. The procedure for separation of phases by centrifugation.

3.3. Phase diagrams: making standards and mixed gels Duplicate standards for microscopy were made from aqueous dispersions of labeled polymer. Concentrations were determined using the micro-assays described below. Desired concentrations were obtained by evaporation in a convection oven at 50–60⬚C. Calibration points in the concentration ranges between 0–4% (pectin) and 0–10% (gelatin) were employed. The linear correlation between fluorescence intensity and concentration was higher for pectin (average R2 ˆ 0:982† than for gelatin (average R2 ˆ 0:848†: Phase-separated mixes of labeled materials for microscopy were made by adding an aqueous solution of sodium chloride to the stirred polymer mix above the gelation temperature of ⬃30⬚C. The desired final concentrations (including 1 M sodium chloride) were obtained by evaporating the mix as described above. The mix was held covered at 60⬚C for additional 0.5–3 h to allow the separation to proceed further. During the quantitative study, samples were placed between horizontal, parallel, pre-heated cover

glasses (thickness #1, rectangular bottom 35 × 50 mm2 ; circular top 25 mm). Standards and mixed gels were prepared under red light to prevent photobleaching. The sample size was 2–5 ml. The mix compositions (unlabeled pectin and gelatin, respectively, per 2.5 ml of 1 M aq. sodium chloride) employed for the construction of quasi-ternary phase diagram from chemical assay were (mix ‘a’:) 0.75%/5.3%, (mix ‘b’:) 1.25%/5.7%, (mix ‘c’:) 1.75%/6%, and (mix ‘d’:) 4.12%/4.6%. 3.4. Centrifuged gel mixes Procedure when making gels for chemical assay: See Fig. 3. 3.5. Micro-assays The ISO hydroxyproline assay (ISO, 1994) based on the approach by Stegemann and Stalder (1967) was modified for the analysis of microgram quantities of gelatin as trans4-hydroxy-l-proline. Modifications included the withdrawal of only 10 ml sample solution, hydrolysis for 24 h in a bath, elimination of the filtration step, and neutralization of the added acid. A potential problem was that the absorption peaks of the assay chromogen and the fluorochrome are very close (555 and 562 nm, respectively). Separate experiments showed that the fluorochrome is chemically destroyed under assay conditions before the absorption is recorded and, thus, no interference takes place. While paying attention to the stability of the reagents, we found our assay well reproducible and relied on one calibration graph during all experiments. The coefficient of variation for any determination of gelatin samples was 1.7%. We have also successfully used this assay for the assessment of collagen in egg shell membranes. The micro-assay, which was linear up to 600 mg gelatin, should, with a changed dilution factor, permit a determination of less than 1 mg gelatin. The method by Scott (1979) for determination of polygalacturonic acid was adopted by AOAC in 1995 as a part of the Official First Action analysis of total dietary fiber (Theander, Aman, Westerlund, Andersson, & Pettersson, 1995). We modified this assay to allow a quantitation limit of 30 mg LM pectin as d-galacturonic acid residues. Modifications included the withdrawal of only 20 ml sample solution, use of cold and relatively less acid, pre-heating of the acidified sample before hydrolysis for 70 min, and elimination of the filtration step. We found this micro-assay to be linear up to at least 900 mg pectin and well reproducible. The coefficient of variation for any determination of pectin samples was 1.3%. 4. Results and discussion The procedure for fluorescent labeling of pectin described in Section 3 should function for all carbohydrate polymers that contain vicinal diols (cis or trans isomers) or when such

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

Fig. 4. Phase separation in a gelatin–LM pectin gel as shown by fluorescent labeling. The sample is illuminated in an ordinary fluorescence microscope with excitation light for the fluorochrome attached to gelatin (left) and pectin (right). The size of each image is 67 × 70 mm2 :

groups can be introduced into the molecule. The first step is the oxidation with periodate (Guthrie, 1962; Jackson, 1944), which has been employed for the oxidation of, for instance, corn- and potato-starch (Jackson, 1944). The periodic acid reaction has been used in combination with the Schiff’s reagent, but histochemical and molecular modeling studies have shown that this combined reaction (called the PAS reaction) is not a quantitative test for polysaccharides (Puchtler, Meloan, & Brewton, 1974). We hypothesized that the use of a bright, less bulky reagent with a spacer and carefully controlled reaction conditions would allow quantitative results. The protocol discussed in this paper results in a 6-atom spacer being located between the fluorochrome and the polysaccharide chain, whereby both steric constraint and transfer of light energy to the chain should be considerably reduced. Katayama et al. (1998) mention that hydrochloric and trifluoroacetic acid have been used as catalysts for the conjugation of hydrazide and steroids in methanol or ethanol solution. However, the use of DMF followed by stabilization of the bonds by treatment with cyanoborohydride resulted in very bright labeling of the pectin without the employment of catalysts. The use of methanol, without catalyst, in the quantitative protocol resulted in a lower but acceptable extent of labeling for quantitative purposes. Commercial DMF was found to have an amine-containing contaminant that slowly reacted with the hydrazide-containing probe to form a dark brown compound. Thus, solutions of BODIPY in DMF should be made fresh for each experiment. Heat treatment in excess of 94⬚C for 1–2 min. was avoided during the making of fluorescent gels, since the BODIPY probe may otherwise not remain stable. The labeling of gelatin and the preparation of samples for microscopy was performed at elevated temperature to avoid gelation and lower the viscosity to facilitate phase separation and handling of the gelatin solutions. A raised temperature was also necessary for phase separation to occur since the phase diagram is inverted with a lower critical solution temperature above the gelation temperature of gelatin (Antonov et al., 1996; Tolstoguzov, 1990). Environmental temperatures in the range 50–60⬚C satisfied the liquefaction, phase separation, and viscosity requirements. Although

583

the solvent DMF tends to precipitate gelatin out of aqueous solutions, no such problems were observed. The use of only two SEC columns in the labeling protocol appeared adequate, since only a very small amount of fluorescent probe remained after the first column. Most of the TAMRA probe could not be eluted from the columns, implying that the probe reacted with the column packing. In addition, gelatin yields lower than 100% indicated that some gelatin was trapped in the column. The procedure for fluorescent labeling of gelatin was, as an alternative, also carried out using a derivative (T-6105) of TAMRA SE with a 7-atom aminohexanoyl spacer between the fluorophore and the reactive group. However, the spacer-equipped fluorophore offered no advantages compared to the standard fluorophore in terms of brightness, gelation, or stability. In a qualitative study, sodium chloride was added to an aqueous mix composed of 1.7% labeled pectin and 9.6% labeled gelatin so that its concentration was 2 M. The mix was held at 55⬚C for 30 min. The fact that sodium chloride actually induced phase separation in a mix of homogeneously labeled gelatin and pectin was demonstrated by withdrawing samples for fluorescence microscopy before and after the addition of sodium chloride. In the first case, only diffuse light emerged from the sample and no contrasts were observed upon excitation of the BODIPY fluorophore. In the second case, complementary light and dark regions were observed when the sample was illuminated with excitation light for each of the two fluorophores. Thus, the light regions represented assemblies of labeled pectin and gelatin rich material, respectively. Both ordinary fluorescence microscopy and CSLM revealed the presence of complementary regions of fluorescence upon excitation of each of the two fluorochromes with suitable light. Digital images of a double-labeled gel are shown in Fig. 4. Each image is the negative of the other, since the same area was illuminated with either blue or green light aimed for BODIPY and TAMRA, respectively. The displayed morphology should be the result of both an initial demixing and a subsequent mixing process, which was halted by gelation. Confocal microscopy was used to obtain a color representation of the distribution of fluorescent labels (Fig. 5). The colors were separated and the intensity values of each color quantified. In the continuous phase the concentration of fluorophore attached to pectin varied more than the concentration of fluorophore attached to gelatin (Fig. 6). This variation, which was on a scale similar to the resolution limit of the instrument, suggests that individual fluorophores were detected, since the labeling of the pectin was sparse. It may not be excluded that clusters of fluorophores were present and detected as peaks. The relatively low apparent fluorescence signal from the gelatin label in the pectin-rich phase may be due to the sigmoidal response curve of the photo-multiplier tube (PMT)-detector of the CSLM equipment. Differences in fluorescence intensity might also be related to uneven illumination within the sample or to

584

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

Fig. 5. Phase separation in a gelatin–LM pectin gel as viewed in the confocal scanning laser microscope. The thickness of the optical slice is about 0.5 mm. The image size is 70 × 70 mm2 : The colors represent: green: pectin label; red: gelatin label; yellow: intermediate composition.

uneven detector response. The CSLM technique permits an estimate of the maximum size of the dispersed particles when the slice thickness is small relative to the diameter of the particle. In the current case, dispersed phase diameters were ⬃20 mm or (usually) less. Fig. 6 shows that the thickness of the interphase region can be estimated at ⬃3 mm. The curvature of the phase boundary will introduce little error (⬍1%) when the thickness to diameter ratio is 0.5:20 as in this case. A portion of the phase-separated mix from which the previous samples had been taken was centrifuged at

20,000g and 40⬚C for 10 min and then immediately cooled to 5⬚C. A small, buff colored phase at the bottom and a sizable, reddish phase of lower density were found in the centrifuge tube. The latter phase was the gelatin rich phase, since only the fluorochrome attached to gelatin is red. According to the phase diagram (Fig. 7) and the results from fluorescence microscopy, the gelatin-rich phase should occupy the largest volume as indeed it did. Previously, while using centrifugation, an unlabeled mix of the same composition had been shown to phase separate with a similar proportion between the phase volumes. Thus, no

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

influence of the labeling itself on the phase separation behavior was observed. Preceding a quantitative study, we determined that a 10% solution of gelatin B at pH 5.5 was not precipitated by the addition of 1 M sodium chloride. This was done because sodium chloride at a concentration of 2 M was found to precipitate gelatin in a 0.56% gelatin solution (Finch & Jobling, 1977). The quantitative study included centrifugation and fluorometry, and, in both cases, micro-assays. The chemical analysis of the different phases in the four centrifuged mixes revealed a typical segregative (i.e. representing polymers of low thermodynamic compatibility) phase diagram (Fig. 7). This is expected, since at pH 5.5 both polymers are similarly charged polyelectrolytes, and the phase behavior resembles that of an aqueous mixture of non-ionic polymers. The analysis also disclosed the occurrence of a three-phase region at high polymer concentrations. The phase-separated region occupies most space in the diagram especially at low solvent concentrations, and this reflects the high molecular weights of pectin and gelatin compared with the molecular sizes associated with the solvent. The apex of the binodal curve and the whole twophase area are located closer to the gelatin axis. This is consistent with a lower solvent compatibility (x pr–s) for gelatin than for LM pectin (x ps–s) and can be regarded as a Dx -effect (Grinberg & Tolstoguzov, 1997). The value of x pr–s for gelatin and water has been estimated to ⬃0.46 (Ziegler, 1988; Ross-Murphy, 1995), and the x ps–s for pectin and water should be slightly lower. The asymmetry of the diagram could also be related to a higher molecular weight

A 240 Green Va lue

220 200 180 160 140 120 97

105

89

81

73

65

57

49

41

33

25

17

9

1

100

Distance

B

250 200 Red Va lue

150 100 50

105

97

89

81

73

65

57

49

41

33

25

17

9

0 1

585

Distance

Fig. 6. Fluorescence intensity of (A) BODIPY FL and (B) TAMRA versus distance across the interphase region in Fig. 5 (1 mm ˆ 7:5 distance units).

AQUEOUS SOLVENT

Threshold point slightly above critical point

2

2

4

4

6 8 % GELATIN 10 12 14 16 18

• a • • ¡b •• •

•

¡¡

c

d

¡

¡

Bulk composition of mix

•

Composition of phase

6 8

% PECTIN 10

• 12

•C

14 16 18

Fig. 7. Ternary phase diagram of an aqueous gelatin–LM pectin system with 1 M sodium chloride and at 60⬚C obtained by using centrifugation and chemical assays.

586

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

of pectin than of gelatin. The tie-lines are slightly skewed towards the gelatin axis, and the critical point (the locus of the tie-lines) is separated from the phase separation threshold point. This is generally expected when xpr–s ⬎ xps–s ; implying a higher water-binding capacity of pectin than of gelatin. The location of the threshold point in a gelatin– pectin mix was determined by Clewlow, Clark, Rowe, and Tombs. (1995a) to be 3.85% gelatin and 0.4% pectin. Our threshold point is located at approximately 4.15% gelatin and 0.9% pectin. The phase diagram is in other respects consistent with results presented by Clewlow et al. (1995a), who determined the phase behavior of a gelatin– pectin mix at 80⬚C, pH 5.5, and 0.5 M sodium chloride concentration. Addition of salt in excess of the amount required for phase separation is not expected to have influenced the phase diagram significantly, but the rate of separation is likely to increase when the concentration of salt is raised. This behavior with respect to salt has been observed on aqueous gelatin–oligosaccharide mixtures (Vinches, Parker, & Reed, 1997). Some depolymerization may take place during the heat treatments of pectin and gelatin, but the effects on the diagram are likely to be small (Clewlow et al., 1995a). In this work, the pectin-rich phase was generally found to be the dispersed phase. This was likely due to use of LM instead of HM pectin and to higher gelatin concentration when compared with the work of DeMars (1995). LM pectin is reported to be less hydrophilic than HM pectin (Walter, 1991) and, thus, could have a higher x ps–s-value, vis-a-vis HM pectin. This would increase the symmetry in the binodal, reduce the width but raise the threshold point of the phase separated area, and likely move the point of phase inversion (Fig. 7). When Dx becomes smaller, the slope of the tie-lines are expected to decrease. It is reasonable to assume that a decreased hydrophilicity, vis-a-vis HM pectin, would make the LM pectin-rich phase less likely to form a continuous phase in aqueous medium. This would be consistent with a shorter distance between the point of phase inversion and the right branch of the binodal. Thus, substitution of LM pectin for HM pectin would make the gelatin-rich phase more likely to be continuous. If the bulk composition point is close to the point of phase inversion, a change in the relative positions of these points will easily take place after such a substitution. The location of the phase inversion point is known to vary considerably with the type of system. Gelatin–agarose and gelatin–maltodextrin gels have phase inversion points at 0.6% agarose/4% gelatin (Tolstoguzov, 1995) and 15% maltodextrin/5% gelatin (Kasapis, Morris, Norton, & Brown, 1993a), respectively. It is reasonable to assume that this variation is related to the higher molecular weight of agarose compared to the molecular weight of maltodextrin. In practice, the location of the point and the solvent partitioning are kinetically controlled and therefore vary with the thermal treatment. For instance, both cooling rate and final temperature influence the location of the phase-inversion point in a

gelatin–maltodextrin system (Alevisopoulos, Kasapis, & Abeysekera, 1996). At approximately 60% sugar, DeMars (1995) found gelatin (4.5%) to be dispersed in HM-pectin (0.5%). However, at 6.0% gelatin and 0.5% pectin concentrations rheological analysis indicated that phase inversion had occurred and gelatin was the continuous phase. In the current case, and in some regions of the sample, none of the two phases seemed to have an apparent dominance. This is consistent with DeMars’ finding. Thus, both the current research and the work by DeMars support that a relatively high gelatin concentration (6–9%) is essential for gelatin to be the continuous phase when the pectin concentration is 0.5–1.5%. The potential for microfluorometry to replace the centrifugal technique and determine the phase diagram in situ was explored by using confocal and ordinary wide-field equipment. There were several reasons for this approach. The latter equipment is less expensive but may still allow attainment of equally good results. The presence of a pinhole in a confocal system limits the light gathering capacity. The contrast and, thus, the resolution that can be obtained in practice may therefore be affected by the chosen pinhole size. In fact, confocal microscopy, which can provide superior image quality in many instances, may produce results that are inferior to ordinary microscopy when sensitivity is a major concern (Stelzer, 1998). The presence of detectors with different sensitivities and ranges in the two systems further motivated a comparative study. In the first case, a confocal scanning microscope with argon- and helium– neon-lasers, and state-of-the-art PMT was used. Such equipment was also used by Kumar, Laird, Srikant, Escher and Patel (1997) in quantitative double-labeling experiments using FITC and rhodamine as fluorescent dyes. In the second case, an inverted fluorescence microscope and a liquid-cooled CCD-camera were employed. Methanol was used to solubilize the fluorescent dye aimed for pectin because of poor stability over time of the dye dissolved in DMF. This change resulted in a more sparse derivatization of the pectin and lower fluorescence intensity. However, this level of derivatization was sufficient because of the higher sensitivity of the CCD detector. Furthermore, a sparse derivatization of the polymers helped avoid self-quenching and possible effects on the phase separation behavior at high concentrations. The observed fluorescence intensity was within the dynamic range of the CCD-camera. In order to avoid fluorescence resonance energy transfer (FRET) in quantitative double-labeling experiments, only one of the polymers in a phase-separated sample was fluorescently labeled. The effects of photobleaching and sample age on the fluorescence intensity were considered. New concentration calibration data for the gelled, labeled polymer were recorded in each experiment. Previous attempts to obtain homogeneously distributed concentrations by evaporation of the polymer solutions on the microscopy slides had not been successful enough. The methodology outlined here allowed two tie-lines to be

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

587

Fig. 8. Phases of labeled pectin 50 min after initialized phase separation in a mix of 1.7% LM pectin, 4.25% gelatin B, and 1 M sodium chloride. The image size is 67 × 45 mm2 :

determined while employing phase separation conditions that closely resembled those previously used during the centrifugation of polymer mixes. The results from microfluorometry are compared with the results based on centrifugation and chemical assays and with literature data. The mix compositions 1.7% LM pectin/4.25% gelatin B and 2.0% LM pectin/5.0% gelatin B were chosen for microscopy. Based on previous results, it was estimated that both these compositions would allow the gelatin rich phase to be the continuous phase. Therefore, all samples would be in the gel state during microscopy at ambient temperature. Pectin only was labeled in two experiments, in which the calibration gels contained between 0.4 and 4% pectin. Gelatin only was labeled in two other experiments, in which the calibration gels contained between 2 and 10% gelatin. In both cases, several samples with holding times varying from 0.5 to 3 h at the temperature of phase separation were studied. The dynamic range of the CCD camera response covered all fluorescence data, although a few low intensities were located close to the non-linear part of the response curve. Photobleaching of the fluorochromes occurred but

Fig. 9. Coalescence of unlabeled pectin phases 50 min after initialized phase separation in a mix of 1.7% LM pectin, 4.25% gelatin B, and 1 M sodium chloride. The image size is 67 × 85 mm2 :

at low rates especially in the case of TAMRA. Accordingly, no correction of intensity data was deemed necessary. Apart from photobleaching, there was an initial decline in fluorescence followed by a significant decrease over a period of several weeks despite that the gel state was maintained. Thus, all measurements of fluorescence intensity were performed within 1–3 days after the initial decline. For any focal depth, the range of light collection was of a magnitude similar to or larger than the thickness of the gel, which was 6–10 mm. Thus, focal setting and minor moisture evaporation from the sample were less critical. Fluorescence intensities were preferentially recorded in the central region of the circular sample and from phases with dimensions at least as large as the gel thickness. Examples of gel morphology are shown in Figs. 8–10. An early stage of separation (i.e. a holding time of ⬃50 min) of pectin-rich phases is shown in Fig. 8. Beginning coalescence of pectin-rich phases is shown as a negative image in Fig. 9, since only gelatin is labeled. Fig. 10 shows a later stage of separation and clustering of pectin rich material when only gelatin is labeled. It also displays that the composition of the gel is likely to be close to the point of phase inversion. Table 1 shows the results from the fluorometric quantitation of pectin and gelatin concentrations in the phases appearing in the thin gels. Comparative data from the centrifugation and chemical assays of bulk mix (Fig. 7) are also displayed. Since it was not possible to expose (all parts of) the samples for centrifugation and fluorometry to identical thermal histories, the results of the two methods are not necessarily identical. Fluorometry and centrifugation gave similar results in the case of labeled pectin. The agreement between the two methods in the case of labeled gelatin is also fair with the exception of the gelatin content of the dispersed phase in the first mix. We believe that a slight non-linearity in the calibration curve could explain this deviation. Generally, the determination of pectin concentrations were associated with smaller statistical error than the determination of gelatin concentrations. Consistent

588

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

when the polymer concentration rises. Thus, there is likely a limit for how high concentrations one can determine, and this limit is expected to be different for each system of mixed food polymers. 5. Conclusions

Fig. 10. Coalesced phases of unlabeled pectin 100 min after initialized phase separation in a mix of 2.0% LM pectin, 5.0% gelatin B, and 1 M sodium chloride. The image size is 280 × 350 mm2:

with the lower correlation among the calibration standards between fluorescence intensity and concentration of labeled gelatin compared to labeled pectin, there could be a more nonlinear behavior at higher concentrations. We believe that these facts are related to a more pronounced associative behavior among molecules of labeled gelatin than among molecules of labeled pectin in aqueous solution. Such phenomena would particularly influence those regions where aggregation of polymer molecules takes place before the start of gelation. Association of chromophore-containing material is likely to occur since water-soluble polymers with hydrophobic groups are known to form microdomains above a critical aggregation concentration (CAC) (Fischer et al., 1998), and clusters or micelles in aqueous solution (Winnik & Winnik, 1993). For instance, proteins conjugated with tetramethylrhodamine are prone to aggregation (Bioprobes 27, Molecular Probes, Inc., OR). This fact suggests that local gelation may take place at a higher temperature than the ordinary gelation temperature of gelatin and freeze in early morphological stages. Thus, variability in observed morphology within samples could be explained on such premises. Although the sparse labeling of the polymers is likely to reduce the influence of the fluorochromes on chain conformation and intramolecular selfquenching, there are increased opportunities for competitive mechanisms by which loss of excitation light energy can occur

Both the centrifugation and fluorometric methods are capable of yielding results of good accuracy when applied to gelled systems. The former method does not include chemical treatment of the polymers, but it includes a centrifugation step and a manual removal of the phases. The centrifugal separation of the phases may not have gone to completion especially if the difference in density between the phases is small or the viscosity of the mix is high. Manual removal of phases is associated with material losses and separation error. These problems are absent when fluorometry is used. In the latter case, the thermal history of the small sample is easily controlled and the integrity of the sample can be preserved during storage. Morphology and kinetic effects can therefore be studied in contrast to when centrifugation is used. Fluorescently labeled polymer may be stored in dry form. The prospect of employing this technique in higher concentration ranges remains and the accuracy of the results will depend on the properties of the particular polymer system. Some further development would be required regarding the preparation of representative slides for fluorometric calibration. Viable analytical approaches based on either ordinary fluorescence microscopy or CSLM can most likely be designed for the determination of the biopolymer concentrations. In the first case, the use of thin samples and a CCDcamera offers the advantages of low-impact fluorescent labeling, minor interfering optical effects, and low cost. In the second case, a confocal scanning microscope with a detector in photon-counting mode could be employed for whole-scanning of bulk samples. An advantage in this case would be a less laborious collection and interpretation of data, if interfering effects can be kept small. Acknowledgements We are grateful to Dr Simon Gilroy and the Department of Biology for advice and providing opportunities for the use of their equipment for fluorescence microscopy.

Table 1 Phase composition data obtained either from fluorometry or from chemical assay. The mixes contained low-methoxyl pectin (P), gelatin B (G), 1.0 M sodium chloride, and water Mix

Phase

% Pectin (Fig. 7)

% Pectin (fluorometry)

% Gelatin (Fig. 7)

% Gelatin (fluorometry)

1.7% P/4.25% G

Dispersed Continuous

2.4 0.5

2.3 0.7

3.5 5.5

2.2 5.1

2.0% P/5.0% G

Dispersed Continuous

3.5 0.5

3.6 0.8

3.4 7.0

3.6 7.5

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

References Alevisopoulos, S., Kasapis, S., & Abeysekera, R. (1996). Formation of kinetically trapped gels in the maltodextrin–gelatin system. Carbohydrate Research, 293, 79–99. Al-Ruqaie, I. M., Kasapis, S., & Abeysekera, R. (1997). Structural properties of gelatin–pectin gels. Part II: effect of sucrose/glucose syrup. Carbohydrate Polymers, 34, 309–321. Aman, P., & Westerlund, E. (1996). Cell wall polysaccharides: structural, chemical, and analytical aspects. In A. -C. Eliasson, Carbohydrates in food (pp. 207–208). New York: Marcel Dekker. Antonov, Y. A., Lashko, N. P., Glotova, Y. K., Malovikova, A., & Markovich, O. (1996). Effect of the structural features of pectins and alginates on their thermodynamic compatibility with gelatin in aqueous media. Food Hydrocolloids, 10 (1), 1–9. BeMiller, J. N. (1996). Gums/Hydrocolloids: analytical aspects. In A. -C. Eliasson, Carbohydrates in food (pp. 265–281). New York: Marcel Dekker. Berth, G. (1988). Studies on the heterogeneity of citrus pectin by gel permeation chromatography on Sepharose 2 B/Sepharose 4 B. Carbohydrate Polymers, 8, 105–117. Blonk, J. C. G., & van Aalst, H. (1993). Confocal scanning light microscopy in food research. Food Research International, 26 (4), 297–311. Blonk, J. C. G., van Eendenburg, J., Koning, M. M. G., Weisenborn, P. C. M., & Winkel, C. (1995). A new CSLM-based method for determination of the phase behaviour of aqueous mixtures of biopolymers. Carbohydrate Polymers, 28, 287–295. Brossmer, R., & Gross, H. J. (1994). Fluorescent and photoactivatable sialic acids. Methods in enzymology, Vol. 247. New York: Academic Press (pp. 177–193). Brunner, A., Minamitake, Y., & Gopferich, A. (1998). Labeling peptides with fluorescent probes for incorporation into degradable polymers. European Journal of Pharmaceutics and Biopharmaceutics, 45 (3), 265–273. Clewlow, A. C., Clark, A. H., Rowe, A. J., & Tombs, M. P. (1995a). Predicting the phase diagram for a gelatin–pectin system. Biochemical Society Transactions, 23, 498S. Clewlow, A. C., Rowe, A. J., & Tombs, M. P. (1995b). Pectin–gelatin phase separation: the influence of polydispersity. In S. E. Harding, S. E. Hill & J. R. Mitchell, Biopolymer mixtures (pp. 173–191). Nottingham: Nottingham University Press. da Silva, J. A. L., & Goncalves, M. P. (1994). Rheological study into the ageing process of high methoxyl pectin/sucrose aqueous gels. Carbohydrate Polymers, 24, 235–245. DeMars, L. L. (1995). Texture and structure of gelatin/pectin-based gummy confections. MSc thesis, The Pennsylvania State University. Dickinson, E., & McClements, D. J. (1995). Advances in food colloids, New York: Blackie Academic and Professional (pp. 11–12). Doner, L. W., & Douds, D. D. (1995). Purification of commercial gellan to monovalent cation salts results in acute modification of solution and gel-forming properties. Carbohydrate Research, 273, 225–233. Durrani, M., Prystupa, D., Donald, A., & Clark, A. H. (1993). Phase diagram of mixtures of polymers in aqueous solution using Fourier transform infrared spectroscopy. Macromolecules, 26, 981–987. EY-Laboratories. (1990). Biotechs. San Mateo, CA: EY Laboratories. Finch, C. A., & Jobling, A. (1977). The physical properties of gelatin. In A. G. Ward & A. Courts, The science and technology of gelatin (pp. 249– 294). New York: Academic Press. Fischer, A., Houzelle, M. C., Hubert, P., Axelos, M. A. V., GeoffroyChapotot, C., Carre´, M. C., Viriot, M. L., & Dellacherie, E. (1998). Detection of intramolecular associations in hydrophobically modified pectin derivatives using fluorescent probes. Langmuir, 14 (16), 4482– 4488. Fulcher, R. G., Faubion, J. M., Ruan, R., & Miller, S. S. (1994). Quantitative microscopy in carbohydrate analysis. Carbohydrate Polymers, 25, 285–293. Gahmberg, C. G., & Tolvanen, M. (1994). Nonmetabolic radiolabeling and

589

tagging of glycoconjugates. Methods in enzymology, Vol. 230. New York: Academic Press (pp. 32–44). Goudsmit, E. M., Matsuura, F., & Blake, D. A. (1984). Substrate specificity of d-galactose oxidase. The Journal of Biological Chemistry, 259 (5), 2875–2878. Grinberg, V. Y., & Tolstoguzov, V. B. (1997). Thermodynamic incompatibility of proteins and polysaccharides in solutions. Food Hydrocolloids, 11 (2), 145–158. Gubenkova, A. A., Somov, V. K., & Shenson, V. A. (1988). Physicochemical properties of pectin and pectin-based solutions and jellies. Pishchevaya Promyshlennost’, USSR, 5, 13–16 (abstract in English). Guthrie, R. D. (1962). Periodate oxidation experimental conditions. In R. L. Whistler & M. L. Wolfrom, Methods in carbohydrate chemistry (pp. 432–435). New York: Academic Press. Haugland, R. P. (1996). Handbook of fluorescent probes and research chemicals, . (6th ed.)Eugene, OR: Molecular Probes (pp. 14, 30). Hermanson, G. T. (1996). Bioconjugate techniques, San Diego: Academic Press. Ichinose, N., Schwedt, G., Schnepel, F. M., & Adachi, K. (1991). Fluorometric analysis in biomedical chemistry, trends and techniques including HPLC applications. Chemical analysis series, Vol. 109. New York: Wiley. Ipsen, R. (1995). Mixed gels made from protein and kappa-carrageenan. Carbohydrate Polymers, 28 (4), 337–339. ISO. (1994). Meat and meat products — determination of hydroxyproline content. ISO 3496:1994(E). Geneve, Switzerland: International Organization for Standardization. Jackson, E. L. (1944). Periodic acid oxidation. In R. Adams, Organic reactions (pp. 341–375). , Vol. 2. New York: Wiley. Kalab, M., Allan-Wojtas, P., & Miller, S. S. (1995). Microscopy and other imaging techniques in food structure analysis. Trends in Food Science and Technology, 6, 177–186. Kasapis, S., Morris, E. R., Norton, I. T., & Brown, R. T. (1993a). Phase equilibria and gelation in gelatin/maltodextrin systems — part III: phase separation in mixed gels. Carbohydrate Polymers, 21, 261–268. Kasapis, S., Morris, E. R., Norton, I. T., & Clark, A. H. (1993b). Phase equilibria and gelation in gelatin/maltodextrin systems — part IV: composition-dependence of mixed-gel moduli. Carbohydrate Polymers, 21, 269–276. Katayama, M., Nakane, R., Matsuda, Y., Kaneko, S., Hara, I., & Sato, H. (1998). Determination of progesterone and 17-hydroxyprogesterone by high performance liquid chromatography after pre-column derivatization with 4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3propiono-hydrazide. Analyst, 123, 2339–2342. Kinsella, J. E., Rector, D. J., & Phillips, L. G. (1994). Physicochemical properties of proteins: texturization via gelation, glass and film formation. In R. Y. Yada, R. L. Jackman & J. L. Smith, Protein structure– function relationships in foods (pp. 1–21). New York: Blackie Academic and Professional. Kumar, U., Laird, D., Srikant, C. B., Escher, E., & Patel, Y. C. (1997). Expression of the five somatostatin receptor (SSTR1-5) subtypes in rat pituitary somatotrophes: quantitative analysis by double-label immunofluorescence confocal microscopy. Endocrinology, 138 (10), 4473– 4474. Mazur, A. W. (1991). Galactose oxidase — selected properties and synthetic applications. ACS symposium series, Vol. 466 (pp. 99–110). Owen, A. J., & Jones, R. A. L. (1998). Rheology of a simultaneously phaseseparating and gelling biopolymer mixture. Macromolecules, 31, 7336– 7339. Ploem, J. S. (1993). Fluorescence microscopy. In W. T. Mason, Fluorescent and luminescent probes for biological activity. Biological techniques (pp. 1–11). New York: Academic Press. Puchtler, H., Meloan, S. N., & Brewton, B. R. (1974). On the binding of Schiff’s reagent in the PAS reaction: effects of molecular configurations in models. Histochemistry, 40, 291–299. Ross-Murphy, B. S. (1994). Physical techniques for the study of food biopolymers New York: Blackie Academic and Professional.

590

T.S. Nordmark, G.R. Ziegler / Food Hydrocolloids 14 (2000) 579–590

Ross-Murphy, S. B. (1995). Small deformation rheological behavior of biopolymer mixtures. In S. E. Harding, S. E. Hill & J. R. Mitchell, Biopolymer mixtures (p. 95). Nottingham: Nottingham University Press. Rost, F. W. D. (1991). Quantitative fluorescence microscopy, Cambridge: Cambridge University Press. Scott, R. W. (1979). Colorimetric determination of hexuronic acids in plant materials. Analytical Chemistry, 51 (7), 936–941. Stegemann, H., & Stalder, K. (1967). Determination of hydroxyproline. Clinica Chimica Acta, 18, 267–273. Stelzer, E. H. K. (1998). Contrast, resolution, pixelation, dynamic range and signal-to-noise ratio: fundamental limits to resolution in fluorescence light microscopy. Journal of Microscopy, 189 (1), 15–24. Strasburg, G. M., & Ludescher, R. D. (1995). Theory and applications of fluorescence spectroscopy in food research. Trends in Food Science and Technology, 6, 69–75. Theander, O., Aman, P., Westerlund, E., Andersson, R., & Pettersson, D. (1995). Total dietary fiber determined as neutral sugar residues, uronic acid residues, and Klason lignin (The Uppsala Method): collaborative study. Journal of AOAC International, 78 (4), 1030–1041. Tolstoguzov, V. B. (1990). Interactions of gelatin with polysaccharides. In G. Phillips, P. A. Williams & D. Wedlock, Gums and stabilisers for the food industry (pp. 157–175). , Vol. 5. Wrexham, UK: IRL Press. Tolstoguzov, V. B. (1995). Some physico-chemical aspects of protein processing in foods. Multicomponent gels. Food Hydrocolloids, 9 (4), 317–332.

Vinches, C., Parker, A., & Reed, W. F. (1997). Phase behavior of aqueous gelatin/oligosaccharide mixtures. Biopolymers, 41 (6), 607–622. Vodovotz, Y., Vittadini, E., Coupland, J., McClements, D. J., & Chinachoti, P. (1996). Bridging the gap: use of confocal microscopy in food research. Food Technology, 50 (6), 74–82. Walter, R. H. (1991). Analytical and graphical methods for pectin. In R. H. Walter, The chemistry and technology of pectin (pp. 207–208). San Diego: Academic Press (pp. 215–216). Walter, R. H., & Sherman, R. M. (1983). The induced stabilization of aqueous pectin dispersions by ethanol. Journal of Food Science, 48, 1235–1237 (see also p. 1241). Wilchek, M., & Bayer, E. A. (1987). Labeling glycoconjugates with hydrazide reagents. Methods in enzymology, Vol. 138. New York: Academic Press (pp. 429–442). Winnik, M. A., & Winnik, F. M. (1993). Fluorescence studies of polymer association in water. In M. W. Urban & C. D. Craver, Structure-property relations in polymers. Spectroscopy and performanceAdvances in chemistry series (pp. 485–505). , Vol. 236. Washington, DC: American Chemical Society. Ziegler, G. R. (1988). Relationship between the dynamic shear modulus of mixed gelatin–egg white gels and their thermodynamic compatibility in an aqueous solvent. PhD thesis, Cornell University. Ziegler, G. R., & Foegeding, E. A. (1990). The gelation of proteins. In J. E. Kinsella, Advances in food and nutrition research (pp. 203–298). Vol. 34. New York: Academic Press.