Comparison of different quantitative biofilm assays
\r\n\tLiterature showed the presence of ACE2 receptors on the membrane of erythrocyte or red blood cell (RBC), indicating that erythrocyte (RBC) can be considered as a peripheral biomarker for SARS-C0V2 infection.
\r\n\r\n\tIncreased levels of glycolysis and fragmentation of RBC membrane proteins were observed in the SARS-C0V2 infected patients, demonstrating that not only RBC’s metabolism and proteome but its membrane lipidome could be influenced by SARS-C0V2 infection changing the homeostasis of the infected erythrocyte. This altered RBC may result in the clot and thrombus formation; the major signs of critically ill Covid-19 patients.
\r\n\r\n\tThis book is going to be a succinct source of knowledge not only for the specialists, researchers, academics and the students in this area but for the general public who are concern about the present situation and are interested in knowing about simple non-invasive measures for identifying viral and bacterial infections through their red blood cells.
",isbn:"978-1-83969-121-8",printIsbn:"978-1-83969-120-1",pdfIsbn:"978-1-83969-122-5",doi:null,price:0,priceEur:0,priceUsd:0,slug:null,numberOfPages:0,isOpenForSubmission:!1,hash:"fa5f4b6ef59e28b6e7c1a739c57c5d2f",bookSignature:"Prof. Kaneez Fatima Shad",publishedDate:null,coverURL:"https://cdn.intechopen.com/books/images_new/10494.jpg",keywords:"Spike Protein, Hemoglobin, Proteins for Oxygen Transport, Altered Protein Structures, RBC ACE Receptors, RBC ACE-2 Receptors, Carboxypeptidase, Mas Receptor, Metabolomics, Gas Transport, Glucose-6-Phosphate, Phosphoglycerate",numberOfDownloads:4,numberOfWosCitations:0,numberOfCrossrefCitations:0,numberOfDimensionsCitations:0,numberOfTotalCitations:0,isAvailableForWebshopOrdering:!0,dateEndFirstStepPublish:"October 15th 2020",dateEndSecondStepPublish:"November 30th 2020",dateEndThirdStepPublish:"January 29th 2021",dateEndFourthStepPublish:"April 19th 2021",dateEndFifthStepPublish:"June 18th 2021",remainingDaysToSecondStep:"3 months",secondStepPassed:!0,currentStepOfPublishingProcess:4,editedByType:null,kuFlag:!1,biosketch:"Dr. Shad is a governing body member and mentor of Women in World Neuroscience (WWN), a division of the International Brain Research Organization (IBRO). She is also a member of IBRO-APRC Global Advocacy responsible for brain research funding distribution in this region.",coeditorOneBiosketch:null,coeditorTwoBiosketch:null,coeditorThreeBiosketch:null,coeditorFourBiosketch:null,coeditorFiveBiosketch:null,editors:[{id:"31988",title:"Prof.",name:"Kaneez",middleName:null,surname:"Fatima Shad",slug:"kaneez-fatima-shad",fullName:"Kaneez Fatima Shad",profilePictureURL:"https://mts.intechopen.com/storage/users/31988/images/system/31988.jpg",biography:"Professor Kaneez Fatima Shad, a neuroscientist with a medical background, received Ph.D. in 1994 from the Faculty of Medicine, UNSW, Australia, followed by a post-doc at the Allegheny University of Health Sciences, Philadelphia, USA. She taught Medical and Biological Sciences in various universities in Australia, the USA, UAE, Bahrain, Pakistan, and Brunei. During this period, she was also engaged in doing research by getting local and international grants (total of over 3.3 million USD) and translating them into products such as a rapid diagnostic test for stroke and other vascular disorders. She published over 60 articles in refereed journals, edited 8 books, and wrote 7 book chapters, presented at 97 international conferences, mentored 34 postgraduate students. Set up a company Shad Diagnostics for the development of cerebrovascular handheld diagnostic tool Stroke meter into a wearable.",institutionString:"University of Technology Sydney",position:null,outsideEditionCount:0,totalCites:0,totalAuthoredChapters:"4",totalChapterViews:"0",totalEditedBooks:"6",institution:{name:"University of Technology Sydney",institutionURL:null,country:{name:"Australia"}}}],coeditorOne:null,coeditorTwo:null,coeditorThree:null,coeditorFour:null,coeditorFive:null,topics:[{id:"16",title:"Medicine",slug:"medicine"}],chapters:[{id:"75447",title:"Detection of Benzo[a]Pyrene Diol Epoxide-DNA Adducts in White Blood Cells of Asphalt Plant Workers in Syria",slug:"detection-of-benzo-a-pyrene-diol-epoxide-dna-adducts-in-white-blood-cells-of-asphalt-plant-workers-i",totalDownloads:4,totalCrossrefCites:0,authors:[null]}],productType:{id:"1",title:"Edited Volume",chapterContentType:"chapter",authoredCaption:"Edited by"},personalPublishingAssistant:{id:"280415",firstName:"Josip",lastName:"Knapic",middleName:null,title:"Mr.",imageUrl:"https://mts.intechopen.com/storage/users/280415/images/8050_n.jpg",email:"josip@intechopen.com",biography:"As an Author Service Manager my responsibilities include monitoring and facilitating all publishing activities for authors and editors. From chapter submission and review, to approval and revision, copy-editing and design, until final publication, I work closely with authors and editors to ensure a simple and easy publishing process. I maintain constant and effective communication with authors, editors and reviewers, which allows for a level of personal support that enables contributors to fully commit and concentrate on the chapters they are writing, editing, or reviewing. I assist authors in the preparation of their full chapter submissions and track important deadlines and ensure they are met. I help to coordinate internal processes such as linguistic review, and monitor the technical aspects of the process. As an ASM I am also involved in the acquisition of editors. 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Biofilms cause severe problems in many natural (Ferris et al., 1989; Nyholm et al., 2002), clinical (Nicolle, 2005; Rice, 2006), and industrial settings (Brink et al., 1994; McLean et al., 2001; Wood et al., 2006), while being beneficial for waste water treatment and biofuel production (Wang and Chen, 2009). In addition, the bioremediation of crude oil spills involves a biofilm of oil degrading microbes, potentially supplemented by marine flagellates and ciliates (Gertler et al., 2010). Identifying the environmental conditions that prevent or support biofilm formation, as well as understanding the regulatory pathways that signal these conditions, is a pre-requisite to both, the solving of biofilm-associated problems and the use for beneficial purposes. In a previous study by our laboratory (Prüβ et al., 2010), it was determined that nutrition ranked among the more important environmental factors affecting biofilm-associated biomass in Escherichia coli K-12. The key to this study was a high-throughput experiment, where biofilm biomass was determined in a collection of cell surface organelle and global regulator mutants under a variety of combinations of environmental conditions. The cell surface organelles each represented a distinct phase of biofilm formation (Sauer et al., 2002). Flagella are required for reversible attachment (phase I), curli or type I fimbriae are characteristic of irreversible attachment (phase II), and a polymeric capsule forms the matrix that permits the maturation of the biofilm (phase III). Eventually, flagellated bacteria are released from the biofilm (phase IV). Phases III and IV are particularly problematic for the disease progression. Bacteria that are located deep within the mature biofilm are particularly resistant to antibiotics and dispersed bacteria tend to serve as a reservoir that continuously feed the infection. Please, see Figure 1 for the distinction of biofilm phases.
The global regulators included in our previous study (Prüβ et al., 2010) are involved in the co-ordinate expression and synthesis of biofilm-associated cell surface organelles. Many of them are components of two-component systems (2CSTS), each consisting of a histidine kinase and a response regulator (for reviews on 2CSTS signaling, please, see Galperin, 2004; Parkinson, 1993; West & Stock, 2001). In response to an environmental stimulus, the sensor kinase uses ATP as a phosphodonor to auto-phosphorylate at a conserved histidine, then transferring the phosphate to the response regulator at a conserved aspartate residue. In addition, many response regulators can be phosphorylated in a kinase independent manner by the activated acetate intermediate acetyl phosphate (for a review on acetyl phosphate as a signaling molecule, please, see Wolfe, 2005). One 2CSTS that is involved in the formation of biofilms is EnvZ/OmpR, regulating the synthesis of flagella (Shin and Park, 1995), type I fimbriae (Oshima et al., 2002), and curli (Jubelin et al., 2005). RcsCDB is involved in the formation of biofilms, serving as an activator of colanic acid production (Gottesman et al., 1985). RcsCDB constitutes a rare phosphorelay, consisting of three proteins and four signaling domains (Appleby et al., 1996). Much of the effect of EnvZ/OmpR, and RcsCDB upon biofilm formation involves FlhD/FlhC (Prüβ et al., 2006), which was initially described as a flagella master regulator (Bartlett et al., 1988) and later recognized as a global regulator of bacterial gene expression (Prüβ & Matsumura, 1996; Prüβ et al., 2001, 2003).
Time course of biofilm formation
An early review article (Prüβ et al., 2006) summarized the portion of the transcriptional network of regulation that centered around FlhD/FlhC. This partial network contained 16 global regulators, among them many 2CSTSs, such as EnvZ/OmpR, RcsCDB, and CpxR. The regulation of approximately 800 genes was affected by the network. Since many of these encoded components of the biofilm-associated cell surface organelles, it was hypothesized that the network may affect biofilm formation. This hypothesis was confirmed by the high-throughput study that led to the identification of nutrition as one of the more instrumental factors in determining biofilm biomass (Prüβ et al., 2010). The global regulators that were part of the network led to the mutant collection for the experiment. Among the tested environmental conditions were temperature, nutrition, inoculation density, and incubation time. Temperature and nutrition were more important in determining biofilm biomass than were inoculation density and incubation time. The mutant screen was consistent with the idea that acetate metabolism may act as a nutritional sensor, relaying information about the environment to the development of biofilms. This hypothesis was confirmed by scanning electron microscopy. A new 2CSTS, DcuS/DcuR, was identified as important in determining the amount of biofilm-associated biomass (Prüβ et al., 2010).
The high-throughput experiment merely determined that nutrient rich bacterial growth media are more supportive of biofilm formation than are nutrient poor media. Specific nutrients that are supportive or inhibitory to biofilm formation were not determined and are the next logical step. This will be dependent on an assay system that quantifies biofilm biomass in the presence of an array of single nutrients. With this study, we will introduce such a system that quantifies biofilm biomass formed by Escherichia coli mutants in the presence of single nutrients by combining the Phenotype MicroArrayTM technology from BioLog (Hayward, CA) with the ATP quantitative biofilm assay that was previously developed by our own lab (Sule et al., 2009), followed up by statistical analysis of the data, and metabolic modeling.
The BioLog Phenotype MicroArray (PM) technology has been developed for the determination of bacterial growth phenotypes (Bochner, 2009; Bochner et al., 2001, 2008). The PM technology consists of 96 well plates with 95 single nutrients dried to the base of each of 95 wells (the additional well constitutes the negative control). When used with the tetrazolium dye that is provided by the manufacturer and indicative of respiration, the PM system is used to determine growth of bacterial strains on single nutrients. Since the total system consists of 20 of such plates, the user is enabled to screen growth under close to 2,000 conditions. The plates are designated PM1 through PM20, with PM1 and PM2 containing carbon sources, PM3 containing nitrogen sources, and PM4 containing sulfur and phosphorous sources. The remaining plates can be used to determine the pH range of growth or resistance to antibiotics or other harsh conditions. Liquid growth media are supplied together with the respective plates.
With respect to bacterial growth, PMs have been used in numerous previous studies (Baba et al., 2008; Edwards et al., 2009; Mascher et al., 2007; Mukherjee et al., 2008; Zhou et al., 2003). However, use of this technology for the investigation of biofilms has been limited (Boehm et al., 2009). In E. coli, the use of PM technology for the quantification of biofilm biomass has not been reported. In addition, the previous use of PM technology in biofilm studies has been based on the use of the crystal violet assay for the quantification of biomass. There are, however, many more assays that have been developed for the quantification of biofilm-associated biomass, each of which serves a different purpose. The different quantitative biofilm assays are compared in Table 1.
Crystal violet is a non-specific protein dye that stains the bacterial cells and their exopolysaccharide matrix for dead and live bacteria alike. Biofilms are cultivated on 96 well plates and stained with 0.1% crystal violet in H2O. In a second step, crystal violet is solubilized with a mix of ethanol and acetone (80:20) and measured spectrophotometrically (O’Toole et al., 1999; Pratt & Kolter, 1998). The assay was developed as a high-throughput assay that is suitable for robotic instrumentation (Kugel et al., 2009; Stafslien et al., 2006, 2007). ATP (adenosine triphosphate) (Sule et al., 2008, 2009) and XTT (4-nitro-5-sulfophenyl-5-[(phenylamino) carbonyl]-2H-tetrazolium hydroxide) (Cerca et al., 2005) are both assays that quantify the energy metabolism of the bacteria. Therefore, only biomass of live bacteria is considered. ATP is converted by the enzyme luciferase into a bioluminescence signal, XTT is reduced by NADH to an orange colored water-soluble formazan derivative. Similar to crystal violet, fluoro-conjugated lectins quantify the biomass of live and dead bacteria alike (Burton et al., 2006). Lectins are highly-specific carbohydrate binding proteins that have been utilized to quantify different cell wall components, as well as extracellular matrix (Stoitsova et al., 2004). Specifically, wheat germ agglutinin (WGA) and soybean agglutinin (SBA) selectively complex lipooligosaccharides and colanic acid, respectively. For our experiments, we needed an assay that quantifies biofilm biomass in live bacteria that is also suitable for high-throughput experimentation, cost effective, and rapid. The ATP assay appeared as the most suitable assay among the five compared assays (Table 1).
Assay | Live/dead cells | Detected material | High-throughput suitability | Reference |
Crystal violet | Live and dead cells | Exopolysaccharide | Yes | (Kugel et al., 2009; Stafslien et al., 2006, 2007) |
ATP | Live cells | Energy (ATP) | Yes | (Sule et al., 2008, 2009) |
XTT | Live cells | Energy (NADH) | Yes | (Cerca et al., 2005) |
WGA | Live and dead cells | Lipooligosaccharide | Not tested | (Burton et al., 2006; Stoitsova et al., 2004) |
SBA | Live and dead cells | Colanic acid | Not tested | (Burton et al., 2006; Stoitsova et al., 2004) |
Comparison of different quantitative biofilm assays
In the past, ATP has been used as a measure of biomass (Monzón et al., 2001; Romanova et al., 2007; Takahashi et al., 2007) because its concentration is relatively constant across many growth conditions (Schneider & Gourse, 2004). For the quantification of biofilms, the BacTiter GloTM assay from Promega (Madison WI) has been used for biomass determination in Pseudomonas aeruginosa (Junker & Clardy, 2007) and E. coli (Sule et al., 2008, 2009). In E. coli, we established that a two fold increase in bioluminescence did indeed relate to a two fold increase in the ATP concentration and a 2 fold increase in the number of bacteria (Sule et al., 2008). Across eight isogenic E. coli strains (one parent strain and seven mutants), differences in biofilm biomass that were determined with the ATP assay were paralleled by observations made with scanning electron microscopy (Sule et al., 2009).
The protocol involves the formation of the biofilms on 96 well micro titer plates, incubation at the desired temperature, and washing of the biofilms with phosphate buffered saline (PBS). Special attention is needed to distinguish the pellicle that forms at the air-liquid interface from the biofilm that forms at the bottom of the wells. In particular, the AJW678 derivatives that we are working with form a solid pellicle that covers the entire surface of the culture (Wolfe et al., 2003). For users who like to include the pellicle into their study, the growth medium and the PBS will be pipetted off carefully from each well. Users who wish to discard of the pellicle can flip the entire 96 well plate over and remove the liquid this way. Eventually, 100 µl of BacTiter Glo reagent are added to each well. After 5 min of incubation, bioluminescence is measured.
For this study, we will use the ATP assay to quantify biofilm biomass that forms on the PM1 plate of BioLog’s PM system. The PM1 plate contains 95 single carbon sources in addition to the negative control. Besides the fact that the use of PM technology for the determination of the nutritional requirements of biofilm has not been reported in E. coli yet, the combination of PM technology with the ATP assay is novel. The combination of both, PM technology and ATP assay, together with the statistical analysis and metabolic modeling, enables the rapid screening of thousands of nutrients for their ability to support or inhibit growth and biofilm formation in one experimental setup. The described technique is not only cost-efficient and easy to perform, but also high-throughput in nature, providing valuable insight into the nutritional requirements during biofilm formation.
The bacterial strains used in this study were the E. coli parental strain AJW678, which was characterized as an efficient biofilm former (Kumari et al., 2000) and its isogenic flhD, fliA, fimA, and fimH mutants. The flhD mutant was constructed by P1 transduction, using MC1000 flhD:kan (Malakooti, 1989) as a donor and AJW678 as a recipient. This resulted in strain BP1094. AJW2145 contained a fliA::Tn5 insertion, AJW2063 a fimA::Kn mutation, and AJW2061 a fimH::kn mutation, all in AJW678 (Wolfe et al., 2003). The mutations abolish expression of FlhD/FlhC, FliA, FimA, and FimH, respectively. As a consequence, mutants in flhD and fliA are non-motile, whereas mutants in fimA are lacking the major structural subunit and mutants in fimH the mannose specific adhesive tip of the type I fimbrium. Bacterial strains were stored at -80C in 8% dimethylsulfoxide, plated onto Luria Bertani plates (LB; 1% tryptone, 0.5% yeast extract, 0.5% NaCl, 1.5% agar) prior to use, and incubated overnight at 37C. Bacterial strains are summarized in Table 2.
Strain | Relevant genotype | Reference |
AJW678 | thi-1 thr-1(am) leuB6 metF159(am) rpsL136 ΔlacX74 | (Kumari et al., 2000) |
BP1094 | AJW678 flhD::kn | (Prüß et al., 2010) |
AJW2145 | AJW678 fliA::Tn5 | (Wolfe et al., 2003) |
AJW2063 | AJW678 ΔfimA::kn | (Wolfe et al., 2003) |
AJW2061 | AJW678 fimH::kn | (Wolfe et al., 2003) |
Bacterial strains used for this study
For this study, a mutation was needed that would abolish one of the early cell surface organelles that contribute to the biofilm, while still permitting the formation of biofilms. We performed scanning electron microscopy (SEM) to determine the ability of the five bacterial strains (parental strain, flhD mutant, fliA mutant, fimA mutant, fimH mutant) to form biofilms. Biofilms were grown for 38 h at 37oC on glass cover slips with tryptone broth (TB; 1% tryptone, 0.5% NaCl) as a growth medium. Biofilms were fixed in 2.5% glutaraldehyde and prepared for SEM as described (Sule et al., 2009). Images were obtained with a JEOL JSM-6490 LV scanning electron microscopy (SEOL Ltd., Tokyo, Japan) at 3,000 fold magnification. 10 to 15 images were obtained per bacterial strain from at least three independent biological samples. One representative image is shown per bacterial strain.
We used the PM1 plate of the BioLog PM system that contains 95 single carbon sources. When used with the tetrazolium dye that is provided by the manufacturer and indicative of respiration (Bochner et al., 2001), the PM system can be used for measuring growth of bacterial strains on single nutrients. We here describe a protocol for the determination of biofilm amounts (Figure 2).
As recommended by the manufacturer for the determination of growth phenotypes, the bacterial cultures were streaked from LB plates onto R2A plates (to deplete nutrient stores) and incubated at 37C for 48 hours. Bacteria were removed from the plates with a flocked swab (Copan, Murrieta CA), resuspended and then further diluted with IF-0a GN/GP Base (BioLog, Hayward CA) inoculation fluid to an optical density (OD600) of 0.1. Leucine, methionine, threonine and thiamine were added at a final concentration of 20 μg/ml, the redox dye that is used for the determination of growth phenotypes was omitted for biofilm quantification. 100 μl of the inoculum was then dispensed into each of the 96 wells of the PM1 plates. The inoculated plates were wrapped with parafilm to minimize evaporation and incubated at 37C for 48 hours. Biofilm amounts were quantified using the previously described ATP based technique (Sule et al., 2008, 2009). Briefly, the growth medium was carefully aspirated out of each well, minimizing loss of biofilm at the air liquid interface. The biofilms were then washed twice with phosphate buffered saline (PBS) in order to remove any residual media components. The biofilms were air dried and quantified using 100 μl BacTiter Glo™ reagent (Promega, Wisconsin, WI). The biofilms were incubated with the reagent for 10 min at room temperature and the bioluminescence was recorded using a TD 20/20 luminometer from Turner Design (Sunnyvale, CA). The bioluminescence was reported as relative lux units (RLU).
The determination of biofilm amounts in the presence of single nutrients was performed four times for each strain. In addition, growth on these carbon sources was determined in three independent replicate experiments, following the protocol that is described for the determination of growth phenotypes and including the redox dye (Bochner et al., 2001). Carbon sources on which both strains grew to an average OD600 of 0.5 or more were selected for the t-test analysis and carbon sources on which each strain grew to an average OD600 of 0.5 or more were selected for the ANOVA/Duncan analysis of biofilm amounts (see below).
Work flow for the determination of biofilm amounts on PM plates with the ATP assay
Prior to the statistical analysis, the biofilm amounts from each strain were normalized for experiment specific variation; total bioluminescence across each experiment was summed up and the fold variation was calculated, using the lowest experiment as a norm (1 fold). Data points in each experiment were divided by the respective fold variation. The normalized experimental data sets were subjected to two independent types of statistical analysis, all done using SAS software (SAS Institute Inc., 2009). First, we performed Student’s t-test on all those carbon sources on which both strains grew to an average OD600 of 0.5 or more to determine statistically significant differences between the amounts of biofilm that were formed on a given carbon source between the two strains. Since this analysis yielded more carbon sources than we could comprehend on a physiological level, we then analyzed each strain individually and then compared biofilm amounts on individual carbon sources for specific nutrient categories of structurally related carbon sources. For this analysis, the normalized biofilm data from each strain were subjected to separate one way ANOVAs, followed up with Duncan’s multiple range tests. The tests compared the means of the amount of biofilm formed in the presence of each carbon source to all the other carbon sources within each strain. Carbon sources whose mean was different from the means of all the other carbon sources with statistical significance formed their own group in the Duncan’s test. Carbon sources whose mean difference from the other carbon sources was not statistically significant formed overlapping groups.
Performing Duncan’s test on the parent strain, two carbon sources formed groups A and B. Among the remaining carbon sources, we determined those that were structurally related to group A and B carbon sources. This was done after a determination of the respective chemical structures with the Kyoto Encyclopedia of Genes and Genomes (KEGG; Kanehisa & Goto, 2000; KEGG, 2006). Biofilm amounts formed by the flhD mutant were compared to the parent strain for all these carbon sources. In a second analysis, one carbon source formed group A in the Duncan’s test for the flhD mutant. Among the remaining carbon sources, we identified two carbon sources that were structurally related. Biofilm amounts for these three carbon sources were compared between the two strains. For both analyses, data were summarized in a Table (3 and 4).
Metabolic pathways that lead to the degradation of all the carbon sources that are discussed in this study were determined with KEGG. Metabolic intermediates that were common between different pathways were used to construct metabolic maps. Pathways for both strains were combined in Figures 5 and 6.
To determine the ability to form biofilm, electron microscopy was performed with the five strains that were listed in Materials and Methods. Figure 3 depicts one representative illustration of the 10 to 15 images that were obtained per bacterial strain. Most of these strains formed biofilm despite mutations affecting cell surface organelles of either reversible (flagella) or irreversible (type I fimbriae) attachment. The sole exception was the fimH mutant which only showed a small number of scattered bacteria attached across the slide. The fimA mutant exhibited a large number of filamentous appendages. We are currently unable to explain these appendages.
Electron micrographs at 3,000 fold magnification for the AJW678 parent strain, and its isogenic mutants in flhD, fliA, fimA, and fimH
We wanted a strain for the phenotype microarray experiment that was able to form biofilm on complex media, while lacking one of the cell surface organelles. Since the amount of biofilm formed by the flhD mutant was similar to that of the parental strain in the electron micrographs, the flhD mutant was selected for further testing using the PM1 plates. The flhD mutant has as an additional advantage that much of the regulation by FlhD/FlhC has been previously described. This vast amount of information will help us to analyze the complex metabolic data.
Biofilms that formed on the PM1 plates were quantified with the ATP assay and compared between the two strains with the t-test. The analysis did not yield any carbon sources that supported more biofilm in the parent strain than in the mutant. The 25 carbon sources that yielded significantly higher amounts of biofilm in the flhD mutant are demonstrated in Figure 4. Since the carbon sources that supported biofilm formation by the mutant more so than by the parent are numerous, we decided to analyze each strain statistically first and focus the comparison between the strains to specific structural categories of carbon sources. These are designated ‘nutrient categories’ throughout this manuscript.
The normalized data set from the parent strain was subjected to Duncan’s multiple range test. According to this test, the two carbon sources that were the best biofilm supporters for the parent E. coli strain, maltotriose and maltose, formed exclusive groups A and B. Without
Biofilm formation in the parent strain and the flhD mutant were compared using a t-test. The dark shaded bars resemble the parent strain, the lighter bars the mutant. The error bars in the graph indicate the standard deviation. Note that only carbon sources were included in this analysis that supported growth to at least 0.5 OD600 in both strains.
forming its own Duncan group, ribose was the carbon source that supported the smallest amount of biofilm among all carbon sources tested, while still supporting growth. The parent strain also formed good amounts of biofilm on the remaining C6-sugars. Interestingly, the amount of biofilm that formed on maltotriose (trisaccharide of glucose) was roughly three times the amount of biofilm that formed on glucose. The amount of biofilm that formed on maltose (disaccharide of glucose) was about twice the amount that formed on glucose. The C5-sugars xylose and lyxose did not support growth of the parental strain to the cutoff of 0.5 OD600. For all these carbon sources, biofilm amounts formed by the flhD mutant were compared to the parent strain (Table 3). In contrast to the parental strain, the flhD mutant did not grow well on C6-sugars and their oligosaccharides. Unlike the parental strain, the mutant did not grow well on ribose, but grew to the cut off of 0.5 OD600 on lyxose and xylose. Still, the amount of biofilm formed by this strain on C5-sugars was low (<1,000 RLU). An interesting phenomenon was observed for sugar phosphates and sugar acids. Sugar phosphates supported biofilm production by the mutant more so (>1,200 RLU) than for the parent strain (<600 RLU). Likewise, sugar acids were found to be good supporters of biofilm for the flhD mutant strain (1,500 to 2,500 RLU), but not for the parent (500 to 800 RLU). This was even more remarkable, considering the fact that the parental strain (OD600 ~ 1.0) grew better on sugar acids than the flhD mutant (OD600 of 0.2 to 0.8).
Nutrient category | Nutrients | AJW678 flhD mutant | |
Biofilm Amount (RLU) | Biofilm Amount (RLU) | ||
Trisaccharide | Maltotriose | 4,935 | NA* |
Disaccharide | Maltose | 2,928 | NA* |
C6-sugars | Glucose Fructose Mannose Rhamnose | 1,615 1,500 1,745 873 | NA* NA* NA* NA* |
C5-sugars | Ribose Lyxose Xylose | 147 NA NA | NA* 650 544 |
Sugar phosphates | Glucose 6-P Fructose 6-P | 614 338 | 1,722 1,258 |
Sugar acids | D-galacturonic acid D-gluconic acid D-glucuronic acid | 668 532 852 | 2,358 1,679 2,110 |
Biofilm amounts on carbon sources which formed their own Duncan’s grouping for the parent strain and structurally related carbon sources. Columns 1 and 2 indicate the nutrient categories and single carbon sources for which data are included. Columns 3 and 4 represent biofilm amounts for the parent strain and the mutant on carbon sources that permitted growth to more than 0.5 OD600. NA denotes ‘not applicable’, where the strain grew to an OD600 below 0.5.
The amount of biofilm formed on each carbon source by the flhD mutant was quantified and subjected to Duncan’s multiple range test. According to the Duncan’s grouping, the sole carbon source that formed its own group A for the flhD mutant was N-acetyl-D-glucosamine. Structurally related carbon sources that were included in the PM1 plate are D-glucosaminic acid and N-acetyl-β-D-mannosamine. Biofilm amounts formed on these three carbon sources were compared between the two strains (Table 4).
Nutrient category | Nutrients | flhD mutant AJW678 | |
Biofilm Amount (RLU) | Biofilm Amount (RLU) | ||
Sugar amines | N-acetyl-D-glucosamine | 4,911 | 1,285 |
D-glucosaminic acid | 660 | NA | |
N-acetyl-β-D-mannosamine | 1,368 | 559 |
Biofilm amounts on carbon sources which formed their own Duncan’s grouping for the flhD strain and structurally related carbon sources. Columns 1 and 2 indicate the nutrient category and single carbon sources for which data are included. Columns 3 and 4 represent biofilm amounts for the flhD mutant and its parent strain on carbon sources that permitted growth to more than 0.5 OD600. NA denotes ‘not applicable’, where the strain grew to an OD600 below 0.5.
On N-acetyl-D-glucosamine, the flhD mutant (4,900 RLU) formed a significantly larger amount of biofilm than the parent strain (1,300 RLU), while both strains grew to approximately 1 OD600. On D-glucosaminic acid, the parent strain did not grow to the cutoff OD of 0.5. The flhD mutant grew well, but the amount of biofilm biomass was poor (~600 RLU). For N-acetyl-β-D-mannosamine, both strains grew well, the flhD mutant expressed more than twice the ability to form biofilm than its isogenic parent.
Metabolic pathways were drawn for the degradation of all those carbon sources that supported amounts of biofilm larger than 1,000 RLU for one of the tested strains. These are carbon sources of the nutrient categories C6-sugars, sugar phosphates, sugar acids, and sugar amines. C6-sugars all have pathways that feed into the Embden-Meyerhof pathway, sugar phosphates are intermediates of this pathway. As shown in Figure 5, mannose, fructose, and N-acetyl D-glucosamine feed into fructose 6-phosphate. Gluconate, glucuronate, galacturonate, and rhamnose feed into glyceraldehyde 3-phosphate. This leads to the production of acetyl-CoA, acetyl phosphate and acetate (Figure 6).
Metabolic pathways from the top biofilm producing carbon sources for both E. coli strains, feeding into the Embden-Meyerhof pathway.
Altogether, we present an assay that builds upon two previous assays, the PM technology and the ATP assay. Both assays have been used in much different contexts previously. PM plates have been commonly used to discover various bacterial characteristics based on phenotypic changes (Bochner et al., 2008). Studies involving PM plates include the evaluation of the alkaline stress response induced changes in the metabolism of Desulfovibrio vulgaris (Stolyar et al., 2007). PMs have also been used for the identification of bacterial species (Al-Khaldi & Mossoba, 2004). The use of PM technology in biofilm research is
Metabolic pathways from the top biofilm producing carbon sources for both strains to the production of acetate. Carbon sources that are printed in bold were top biofilm supporters for the parent strain. Carbon sources that are underlined were top biofilm supporters for the flhD mutant. The effect of acetyl phosphate on RcsB and OmpR on the synthesis of flagella, curli, fimbriae, and capsule is indicated.
limited to a study of the ability of E. coli to form biofilm upon ribosomal stress (Boehm et al., 2009). That study used the crystal violet assay as a detection tool for the amount of biofilm.
Here we report for the first time a combination of the established ATP assay along with the PM technology to assess nutritional dependence of E. coli during biofilm formation. Since the statistics approach alone (t-test) yielded no more than a list of data that were difficult to interpret, we decided for a combined statistics/metabolism approach to analyze the complex data. The combination of the two experimental parts of the assay together with the two analysis parts enables the user to rapidly screen hundreds and thousands of single nutrients for their ability to inhibit growth and biofilm formation in one experimental setup. Integrating different mutants into the study will yield valuable insight into the regulatory mechanisms that are involved in the signaling of these nutrients. The described technique is not only cost-efficient and easy to perform, but also high-throughput in nature. It is ideally suited to provide valuable insight into the nutritional requirements that determine biofilm biomass, as well as the respective signaling pathways.
In the described study, we observed that the FlhD mutants made quantitatively higher amounts of biofilms on numerous carbon sources. Interestingly, the parental strain did not form higher quantities of biofilm than the mutant on any of the tested carbon sources. These observations shed light into the ongoing controversial debate, elucidating the role of motility in biofilm formation. In certain bacterial species including Yersinia enterocolitica, the presence of motility has been shown to be beneficial for biofilm formation (Wang et al., 2007). Several previous studies from our lab demonstrate that the absence of motility enhances the ability of E. coli to form substantial amounts of biofilm. As one example, strains transformed with the FlhD expressing plasmid pXL27 showed diminished biofilm forming capabilities (Prüß et al., 2010). Additionally, ongoing studies carried out in the lab with E. coli O157:H7 and the E. coli K-12 strains MC1000 and AJW678 point in the same direction, exemplifying our belief that FlhD and motility are detrimental to biofilm formation for our bacterial strains and under the conditions of our experiments (Sule et al., unpublished data).
As a second observation, carbon sources that supported maximal biofilm formation by either strain all fed into glycolysis eventually, and produced actetate. Although the carbon sources that promoted the highest biofilm amounts were different for the two strains, they still were in the same pathway. The previous high-throughput experiment that had pointed towards nutriition as instrumental in determining biofilm associated biomass had also postulated acetate metabolism as one of the key players in biofilm formation (Prüß et al., 2010). Phosphorylation of OmpR and RcsB by the activated acetate intermediate acetyl phosphate (Kenney et al., 1995) and acetylation of RcsB by acetyl-CoA (Thao et al., 2010) have been described in the past. These activated 2CSTS response regulators then affect the expression level of biofilm associated cell surface organelles, such as flagella, type I fimbriae, curli, and capsule (Ferrieres & Clarke, 2003; Francez-Charlot et al., 2003; Oshima et al., 2002; Prüß, 1998; Shin & Park, 1995) (Figure 6). The positive effect on biofilm amounts of carbon sources that lead to the production of acetate can be explained with the combined inhibitory effect of acetyl phosphate and acetyl-CoA on flagella through OmpR and RcsB and the above described disadvantage of flagella and motility during biofilm formation. We however do not state that acetate is the sole controlling mechanism as the complexity of the bacterial system cannot be explained based on a small number of signaling molecules.
The most striking observation obtained from our studies pertains to the pattern of growth and biofilm formation on sugar acids. It was observed that the FlhD mutants grew to lower optical densities on sugar acids, but formed much higher amounts of biofilm as compared to the parental strain. Previous work from the Prüß lab had shown similar defects in growth of flhD mutants on sugar acids (Prüß et al., 2003), biofilm formation was not tested in that study. The inverse effect of sugar acids on growth and biofilm amounts may have implications in the intestine. Mutants in flhD have an early disadvantage in colonization, but recover after prolonged incubation (Horne et al., 2009). They even take over the population after more than two weeks (Leatham et al., 2005). The initial lack of colonization could be explained by the inability of the flhD mutant to degrade the numerous sugar acids present in the intestine (Peekhaus & Conway, 1998). On the other hand, the ability to take over the bacterial population at a later stage may have to do with the lack of the flagellin, which is a potent cytokine inducer (McDermott et al., 2000). The here discovered ability to make an increased amount of biofilm may add to the long term survival of flhD mutants in the intestine. Bacteria deep within the biofilm will be protected from the immune system, while metabolizing very slowly and not needing much nutrition.
Among the carbon sources that were the least supportive of biofilm formation, the inability of the C5-sugars to support growth and/or biofilm formation was the most striking. Ribose supported growth by the parent strain, but yielded the lowest biofilm amount of all tested carbon sources. The flhD mutant did not even grow on ribose. According to Fabich and coworkers (Fabich et al., 2008), ribose is not among the carbon sources that the E. coli K-12 strain MG1655 utilizes when bacteria colonize the intestine. Our data are consistent with this observation. Since E. coli O157:H7 EDL933 does actually utilize ribose in the intestine, ribose utilization may constitute a mechanism by which pathogenic E. coli can find a niche in the intestine to co-exist with the commensal E. coli strains.
The inability to grow on lyxose is also consistent with previous observations, where only a mutation in the rha locus enabled the bacteria to grow on lyxose via the rhamnose pathway (Badia et al., 1991). Normally, E. coli are unable to grow on lyxose. Most interesting is the behavior of the two strains on xylose. The parent E. coli strain was unable to grow on xylose. The flhD mutant did grow, while producing moderately low amounts of biofilm. Co-utilization of glucose and xylose by E. coli strains is of upmost importance during the production of biofuels, since the fermented plant material contains both, cellulose (polymer of glucose) and hemicellulose (polymer of glucose and xylose), in addition to lignin. Much research is currently dedicated to the genetic modification of E. coli that enables the bacteria to utilize xylose more efficiently (Balderas-Hernandez et al., 2010; Hanly & Henson, 2010). It would be interesting to see whether a mixture of our parent strain and its isogenic flhD mutant would be able to co-utilize glucose and xylose, particularly since the mutant produced a moderate amount of biofilm which can also be beneficial to the production of biofuels.
In summary, we developed an assay system that quantifies biofilm biomass in the presence of distinct nutrients. The assay enables the user to screen a large number of such nutrients for their effect on biofilm amounts. Examples of metabolic analysis relate back to previous literature, as well as giving raise to new hypotheses. Yielding further evidence for the previous hypothesis that acetate metabolism was important in determining biofilm amounts can serve as a positive control that the assay actually yields data of biological significance. Particularly with respect to life in the intestine and the production of biofuels, the data open new avenues of research by providing testable hypotheses. Overall, there is no limit to extensions of the assay into different bacterial species or serving the development of high-throughput data mining algorithms that will computerize the statistic/metabolic analysis that we started in this study.
The authors like to thank Dr. Alan J. Wolfe (Loyola University Chicago, Maywood IL) for providing the bacterial strains that were used for this study, Dr. Jayma Moore (Electron Microscopy Lab, NDSU) for help with the scanning electron microscopy, Dr. Barry Bochner (BioLog, Hayward CA) for helpful discussions during the development of the combination assay, and Curt Doetkott (Department of Statistics, NDSU) for performing the statistical analyses of our data and helping us with their interpretation. The work was funded by an earmark grant on Agrosecurity: Disease Surveillance and Public Health through USDA/APHIS and the North Dakota State Board of Agricultural Research and Education. Figure 2 was created using Motifolio (Motifolio Inc., Ellicott MD).
The Republic of North Macedonia is a land-locked country in southeastern Europe in the Balkan Peninsula, with 850 km of frontier with five countries: Serbia, Kosovo, Montenegro, Albania, Bulgaria, and Greece. The country has a surface area of 25,713 km2. Half of this area (1.26 Mio. ha) is agricultural land, out of which 560,000 ha are classified as cultivated land and 704,000 ha as permanent pastures. Mountainous forest land covers 37% of the country, and about 2% is covered by lakes. Livestock and goat farming is very important in the Polog area, in the foot of the Sharr Mountain. These areas represent a mountain chain stretching from the southern part of Kosovo and northwestern Macedonia to northeastern Albania. The Sharr Mountain represents the largest mountain massive in Macedonia and extends to these geographic coordinates, between 42°41′43″ and 42°16′34″ north latitude, as well as between 20°34′51″ and 21o16′00″. The mountain system is about 80 km long and 10–20 kilometers wide. It includes a number of high points, among which the highest peak of the Titov vrv is 2747 m, Mali Turc (2702 m), Ljuboten (2498 m), and Bistra (2641 m).
Goat breeding in Macedonia is defined by spontaneous and continuous development, and with each day, there is major concern of farmers for goat’s growth as a market that provides secure subsistence and business. The breed structure of goats in the country is based on the domestic Balkan goat with a certain representation of the Alpine breed goats, Saanen, and crossbreds of these races. The basic product obtained from goats is goat’s milk (which is commonly processed into white-brined cheese, yogurt, and kashkaval), kids, and goat meat [1]. The official restriction of goat breeding in the territory of Macedonia in 1947 resulted in disastrous consequences, which lasted for more than 40 years. The goats in the entire territory of the former Socialist Republic of Macedonia were slaughtered and rapidly reduced number from 516,800 in 1947 to 47,000 in 1949 or more than 90%. However, in 1989, goat production was again allowed, and since then, the interest of the farmers in goat production has increased [2].
Pacinovski et al. [3] reported that Goat livestock industry has a century-old tradition in Macedonia due to environmental factors and a type of vegetations suitable for goat breeding; wherefore, these animals have provided subsistence of the population in the past centuries. Historically, the number of goats bred in Macedonia was around 500,000 heads, with the law for prohibited goat breeding (Law for Prohibited Goat Breeding, 1948). This law had highly significant negative role in reducing the number of goats in the Republic of Macedonia. Petrovska et al. [4] emphasized that goat breeding in Macedonia is difficult due to a number of factors such as unfavorable racial composition and fragmentation of herds, unorganized and insecure sale of milk and dairy products, shortage of labor, weak and irregular application of selective measures in the herds, and more. However, compared with past years, it can be concluded that there is spontaneous and continuous development of the husbandry industry with growing interest of farmers for goat breeding. The total number of goats the Republic Macedonia is around 80,000 with a tendency to increase.
There are six genotypes (breeds) of goats present in Macedonia in the system for identification and marking of livestock within the Food and Veterinary Agency: domestic Balkan goat, Alpine, Saanen goat, crossbreeds with Alpine, crossbreeds with Saanen goat, and genotype registered under the name of other population. According the same agency, out of the total quantity in 2011, 48% of the total number of goats are domestic Balkan goats, 5.5% are Alpine goats, 7.9% are crossbreeds with Alpine goat, 7.8% are Saanen goats, 3.4% are crossbreeds with Saanen goat, and the rest are recorded as other breeds of goats [5].
According to Pacinovski et al. [6] and the data of the Food and Veterinary Agency, the department for identification and registering of domestic animals, there are six genotypes of goats in the Republic of Macedonia: domestic Balkan goat, Alpine, Saanen, Alpine crossbreed, crossbreeds with Saanen goat, and population registered under the term of other. The most represented goat breed in the country is domestic Balkan goat, with a number of around 38378 goats, goats registered as other with a number of 21772 goats, the number of crossbreeds with Alpine is 6330, Saanen with 6256 goats, Alpine is represented with 4193 and crossbreeds with Saanen are represented with 2735 goats. Balkan goat is well adopted to the existing climate conditions in the country as well as to the existing nutritional resources especially in the hilly mountainous areas of the Republic of Macedonia, which are not suitable for other domestic animals. It is the shrubbery vegetation which is especially attractive to goats. The excellent adaptability of the breed is due to the excellent health condition of goats manifested during the whole year. Compared to the other breeds (Alpine, Saanen, and crossbreeds between the same with other breeds), Balkan goat is extremely resistant to many diseases (chronic, bacterial, etc.). They are especially resistant to emergent climate changes that affect the goat health.
Traditional cheeses represent a cultural heritage and are the result of accumulated empirical knowledge passed from generation to generation [7]. Accurate and precise milk recording is one of the most significant moments for a successful selection of milking goats. In this context, breeders are constantly making efforts to find the most suitable and cheapest methods for conducting of tests for milk. According to Pacinovski et al. [8], in Macedonia predominate extensive goat dairy industry and machine milking are not widespread throughout the country. There is much to be done about the improvement of goat farms in respect to goat breeding and comprehensive mechanization of farm routines which both increase efficiency of the farms.
Milk and cheese samples were analyzed in duplicate for moisture, fat, salt, pH, titratable acidity (as percentage of lactic acid), and total nitrogen [9]. Total nitrogen (TN) content was estimated by the Kjeldahl method using a Kjeldahl device (model DS1; Simsek Laborteknik, Ankara, Turkey) [10]. The water-soluble nitrogen (WSN) and 12% TCA-soluble nitrogen (TCA-SN) as percentage of TN and total free amino acid (FAA) of the cheeses were determined [11].
The water-insoluble fractions of the cheeses were freeze-dried and then analyzed by urea-PAGE using a Protean II XI vertical slab gel unit (Bio-Rad Laboratories Ltd., Watford, UK), and the gels were stained directly with Coomassie Brilliant Blue G-250 dye [12, 13].
After destaining using pure water, gel slabs were digitized using a scanner (HP Scanjet software, Scanjet G4010; Hewlett-Packard, Palo Alto, CA). Scans of the electrophoretograms were used to quantify bands using densitometric software (ImageMaster TotalLab Phoretix 1D Pro software; Keel House, Newcastle upon Tyne, UK).
The caseins and peptides were determined quantitatively by integration of peak volumes and areas using the densitometer. The WSN fractions of the cheeses were also freeze-dried for determination of peptide profiles. The analysis was realized by RP-HPLC using a Shimadzu LC 20 AD Prominence HPLC system (Shimadzu Corp., Kyoto, Japan) [14].
Solid-phase microextraction/GC-MS analysis of volatiles. Analysis of the volatiles was performed by a static solid-phase microextraction method, using aGC-MS system (Shimadzu Corp.). The identifications were based on comparing mass spectra of unknown compounds with those in the mass spectral library of John Wiley and Sons Inc. (2005) and the National Institute of Standards and Technology/Environmental Protection Agency/National Institutes of Health (NIST/EPA/NIH 02;
A total of 33 authentic standard compounds (Sigma Chemical Co., St. Louis, MO) were used to confirm the identities of volatile compounds in the cheese samples. The concentrations were calculated by the comparison of the peak areas of the internal standard containing mixture of 2-methyl-3-heptanone and 2-methyl-1-pentanoic acid in methanol (Sigma-Aldrich Co.) and unknown compounds. Each compound was expressed in micrograms per 100 g of cheese.
Goat breeding is an important livestock branch, and great attention is paid to its development and industrialization in all Mediterranean countries. The composition of milk is of great importance for determining the technological properties of goat milk and for its further processing in a suitable type of cheese. According to the historiographic data, the golden period of the Macedonian livestock breeding was in the middle of the nineteenth century, when there were about 7–9 million sheep and 2 million goats in Macedonia’s borders [8].
With the adoption of the Law on Goat Breeding in 1989, the goat again began to take its place in our livestock breeding. Today’s situation of goat is heterogeneous, given the fact that the individual producers preserve 1–2 goats, but there are also those who breeds 20–30 goats, while the organized goat farms have 100 or more heads. It is estimated that in our country, there are under 100,000 goats with a major participation in the domestic Balkan goat and in a smaller proportion the Saanen, Alpine, and their half-bred [6].
Accordingly, the milk used for the production of cheese should be of a normal chemical composition in accordance with the rulebook on requirements for the quality of raw milk, quality standards for consumer milk, dairy products and the use of their names, quality and activity of starter cultures, curdling and other specific substances, and the manner of their use, the manner of additional labeling of milk and dairy products, as well as the permitted deviation of weight in relation to the declared [15]. Goat milk is often mixed with the cow or sheep’s milk and as such is offered to the dairies so that its nutritional values are not properly valuated. A certain part is processed by farmers in white goat cheese, but due to its specific taste and smell, it consumes a certain number of the population, although it has higher nutritional and therapeutic values compared with other cheeses. Goat breeding has a good perspective, given the good natural conditions, but also because of the fact that goat milk is given greater importance nowadays, especially because of its dietary and nutritional properties.
The quantities of individual ingredients affect the technological characteristics of the curd, as well as the organoleptics and the quality of the finished product [16]. The physical-chemical characteristics of the used milk for the production of the white-brined goat cheese was 3.44 ± 0.10 g/100 g for protein, 3.00 ± 0.24 g/100 g for fat, 11.99 ± 0.17 g/100 g for total solid, and 4.79 ± 0.09 g/100 g for lactose; the total microbiological counts were 5.49 ± 2.119103 log cfu/mL, and the milk total somatic cell numbers were 85 ± 0.109103 cell/mL [10]. The pH of the milk was 6.60. From the obtained analysis, the raw goat milk fulfill the conditions according to the book of rules for hygienic criteria and milk quality [15, 17]. Given that the composition of milk is highly variable and depends on numerous genetic and paragenetic factors, its comparison shows great differences with the findings of other authors (Table 1).
Characteristics | Mean ± SD (n = 30) |
---|---|
Total solids | 12.64 ± 1.240 |
Fat | 3.84 ± 0.360 |
Protein | 3.21 ± 0.034 |
Casein | 2.49 ± 0.031 |
Lactose | 4.49 ± 0.077 |
Ash | 0.75 ± 0.027 |
pH | 6.65 ± 0.056 |
Kashkaval, white-brined, and beaten cheese are the three main types of cheeses produced presently in Macedonia. The origin of beaten cheese is from the territory of Mariovo, produced in the past years on the pasture land only from ewe’s milk. According to its salty taste and its hard consistency, it is an authentic product with characteristics that is preserved even in usual situations. The production of cheese has been carried out since the time of the Ottoman Empire. The “beaten” designation is originated from the one process step of the cheese production where the cheese curd is beaten to ensure proper draining (Figure 4) [18, 19].
Cilev et al. [5] investigated the chemical composition of goat milk on three farms during the month of April, and the highest percentage of milk fat is determined in milk from a farm in Kožle (3.85%), and the lowest percentage in milk is from a farm in Ajvatovci (3.50%). In terms of protein content, the highest percentage (3.70%) is determined to a farm in Taor, and the lowest is in the farm in Ajvatovci (3.05%). The content of lactose was highest in the farm in Ajvatovci (4.71%), while the lowest is from a farm in Taor (4.43%). The highest content of fat-free dry matter was found on the farm will be displayed (8.69%), while the lowest farm is Taor (8.26%). The total dry matter in milk was also highest in the farm will be displayed (12.42%) and the lowest farm is Taor (11.85%). In terms of the content of added water, the result charter in April in all three farms was zero, which indicated its full functionality in terms of physical water added.
Brined cheeses are with high salt content, which enables their preservation even in the warm periods of the year. They are produced from sheep, goat, and buffalo milk, as well as from their combinations. During the ripening, changes in the composition and properties of the cheeses are mutually dependent on changes in the brine [20, 21, 22].
In the last few years, the increased interest of the goat’s milk products on the marketplace and the scientific community is consistent with the general trend and efforts for the production of healthy food, since the goat’s milk has been well-known for its beneficial effects on human health [23]. According to statistics in 2011, white-brined cheeses are consumed in quantities of 7.4 kg per year, followed by 2.2 kg kashkaval cheese and urda (ricotta) with 2.1 kg by member of households [24].
Sulejmani [25] reported that white-brined cheeses have a high salt content that allows them to stay in the warm periods of the year. They are produced from sheep, goats, buffalo milk, and their combinations. In the ripening, changes in the composition and properties of the cheeses are mutually dependent on changes in the brine. Most varieties in this group are stored in closed containers, but some are stored in gas-permeable containers, which affect biochemical changes that occur in the process of ripening and storage.
The milk for beaten cheese manufacture is drained through cheesecloth (not obligatory) and poured into a curdling vessel. The curdling is most often done using enzymatic rennet with the strength of 1:5000 or the rennet chymosin CHY-MAX (2080 imcv/g) at the temperature of the milk of 25–35°C. In the past, for curdling homemade rennet was used obtained from the lamb’s stomach. The curdling process lasts 30–50 min. After that the curd is submitted to processing (churning or beating) using a wooden tool. The process of churning (beating) is done in 3 series of 50 strokes (150 strokes in total), and after each series, the curd is left to “rest” for 5–10 min. In this process it may come to separation of a part of the milk fat, in which case the fat is skimmed and removed from the vessel. When the beating process ends, the curdled mass is warmed up by adding warm water to the temperature of 53–90°C, depending on the particular manner of production [27] (Figure 1).
Typical beaten cheese production with a mixture of goat milk. After Sulejmani [26].
Recently consumers are more aware about the relationship between their eating habits and nutritional status. Consequently, they look for foods that are added with natural products rather than synthetic chemical compounds. Currently, they have interest in maintaining good health and an excellent body figure; therefore, they have become more careful in the food they choose to consume, looking for food with a high nutritional value, bioactive compounds, and antioxidant capacity, such as herbs, fruits, and vegetables. This is an opportunity for some local producers to manufacture cheese products with the partial or total replacement of those chemical additives by natural herbal not only because of their antioxidant but also antimicrobial properties (Figure 2).
Beaten goat milk cheese with Origanum vulgare.
Antioxidant capacities of beaten goat cheeses, of 7 and 20 days ripened cheese (matured cheese), were higher than beaten cheese without plants (Sulejmani and Hayaloglu, unpublished data). Therefore, it could be hypothesized that consumption of matured white cow cheese could notably contribute to the body’s antioxidant defense and prevention of diseases related to oxidative stress. However, further research is needed to elucidate the role of herbs in the antimicrobial and anticancer protective functions in human. Origanum vulgare is a perennial herbaceous plant, with wood stalk. The stub is usually gray-eyed. The roots are superficial, with a multitude of roots reaching at depths of 3–4 cm, and the plant is easily pulled. The flowers are short-tailed, gathered in a long spike in the midst of strong scent bows. It flourishes from the end of June and continues until the end of August.
Red oregano has reddish flowers and a pleasant smell. The flowers are full of nectar and are always frequented by bees. It is aromatic and spicy medicinal plants. The excellent ethereal oil is extracted from this. From 100 kg of dry matter, 2–2.25 kg of ethereal oil is extracted. Oregano on leaves and flowers contains etheric oils in various quantities consisting of a series of special value components. Essential oil (maximum 4%) may contain variable amounts of phenol, carvacrol, and timol. In addition, there are variants of monoterpenes, hydrocarbons (limonene, terpene, ocimene, caryophyllene, β-bisabolene, and ρ-cymene) as well as alcoholic monoterpene (linalool 4-terpineol). It has important properties, as antioxidants, antibacterial, antifungal, and anti-inflammatory and, recently, as anticancer. Oregano possesses powerful properties like antioxidants comparable to those of ascorbic acid and vitamin E. Carvacrols, thymol, and rosemary acids are the main components of essential oil. Known as a food supplements, carvacrol is a potent and bacteriostatic useful against mold and bacteria.
The influence of different heat treatments on goat milk was studied in detail [16]. Multiple analyses confirmed that the heat treatment of goat milk delays the initial coagulation and syneresis, and it improves the retention of dry matter, fat, and proteins. Therefore, on the basis of this finding, technological approaches of white-brined cheese were developed (Figure 3). The characteristic ability of goat milk proteins to retain water; the specific structure and the rheological properties of the cheese curd enable optimal regulation of the fermentation process of cheese and its salting. Traditionally, this type of cheese has been produced by local farmers on a small scale for decades using raw milk, and traditional techniques handed down from generation to generation using only elementary equipment. Instead of using a commercial starter culture, artisan cheese makers relies on the indigenous natural present microorganisms in the raw non-pasteurized milk and adventitious contaminants from the soil, equipment, surfaces, and the environment in general.
Schematic illustration of white-brined cheese making [16].
Schematic illustration of industrial (1) and traditional (2) beaten cheese production using goat/ewes milk combination [26].
Sulejmani and Hayaloglu [18] investigated the use of raw and pasteurized goat milk in the production of Macedonian white cheese. Milk was collected from a certified organic farm from a Saanen goat’s herd of a Novacani village (Veles, Macedonia). Two batches of cheeses from pasteurized (80°C for 2 min) (GP) and raw (GR) goat milk were produced traditionally using artisanal protocols. Goat milk coagulation was attained with commercial enzyme (1 g/100 per L milk) with a stated power of coagulating from 2235 IMCU/g (Chr. Hansen, Powder Extract CHY-MAX, Hørsholm, Denmark). The milk was coagulated at 32°C for 45 and 120 min for GR and GP cheeses, respectively. The coagulum was cut to medium-size (1–2 cm) grains. After whey removal by pressing, cheeses with block form weighing 0.5–1.0 kg were pressed for 4 and 8 h for GR or GP cheeses, respectively. At last, both cheeses were ripened in brine (15% w/v at 4°C) for 120 days.
The chemical composition of white-brined goat cheese made from pasteurized (GP) or raw (GR) milk at the first day was as follows: pH, 5.25 and 6.27; fat-in-dry matter, 37.86% (w/w) and 43.36% (w/w); dry matter, 32.60% (w/ w) and 33.00% (w/w); fat, 12.38% (w/w) and 14.25% (w/ w); and salt, 2.02% (w/w) and 2.73% (w/w), respectively. The use of pasteurization significantly affected the total solid, fat, moisture, and fat-in-dry matter contents of the cheeses (P < 0.05). The white cheese chemical composition was in compliance with the official bulletin [15]. Higher cheese pH levels were found in the cheeses made from raw milk (GR) compared with pasteurized milk (GP) (P < 0.05).
The values of WSN and 12% TCA-SN (expressed as percentage of TN) of both white goat cheeses are presented in Tables 2 and 3. The quantity of WSN and TCA-SN in the cheeses increased during ripening (until 60 days); however, after that, the increase was not intense during the end of ripening. After 60 days of ripening, GR cheeses had higher quantity of WSN than GP cheeses; also the highest TCA values were recorded at day 60 of ripening and then declined again. However, the quantities of WSN were higher in GR cheeses than in GP cheeses (P < 0.05). Higher and similar quantities of TCA-SN and WSN at 60 days of ripening were found in Teleme white-brined goat cheese, respectively [28].
Parameters | Cheeses | Ripening time (days) | ||
---|---|---|---|---|
1 | 60 | 120 | ||
Total | GR | 10.19 ± 0.95bA | 8.43 ± 0.04aA | 9.59 ± 0.79abA |
Protein | GP | 10.13 ± 0.94aA | 9.90 ± 0.42aB | 10.88 ± 0.20aB |
WSN-SN | GR | 7.74 ± 1.01aA | 8.64 ± 0.37aB | 7.31 ± 1.02aB |
(% of TN) | GP | 5.82 ± 0.37bA | 6.22 ± 1.58bA | 2.93 ± 0.37aB |
TCA-SN | GR | 2.44 ± 0.65aA | 3.05 ± 0.02aB | 2.92 ± 0.46aB |
(% of TN) | GP | 1.71 ± 0.24abA | 2.39 ± 0.52bA | 1.44 ± 0.21aA |
TFAA | GR | 0.41 ± 0.00eB | 0.30 ± 0.01aA | 0.44 ± 0.02dB |
mg Leu/g | GP | 0.35 ± 0.00cA | 0.30 ± 0.00bA | 0.39 ± 0.01dA |
Chemical parameters during ripening in raw (GR) and pasteurized (GP) white-brined goat cheeses.
SD, standard deviation; TFAA, total free amino acid; DM, dry matter; WSN, water-soluble nitrogen. TN, total nitrogen; TCA, 12% trichloroacetic acid-soluble nitrogen. a, dMeans ± SD within a row and A–BMeans ± SD within a column with no common superscript capital letters differ (P < 0.05), respectively. Adapted from Sulejmani and Hayalogu [18]
Parameters | Kumanovo | Radovish |
---|---|---|
pH | 5.01 ± 0.01 | 5.43 ± 0.05 |
% Lactic acid | 1.73 ± 0.10 | 4.45 ± 0.13 |
Dry matter, % | 68.43 ± 0.11 | 57.34 ± 0.21 |
Moisture, % | 31.57 ± 0.11 | 42.67 ± 0.21 |
Fat, % | 29.63 ± 0.25 | 26.00 ± 0.41 |
Fat(dm), % | 43.29 ± 0.35 | 45.35 ± 0.84 |
Salt, % | 4.11 ± 0.07 | 8.18 ± 0.38 |
Proteins, % | 32.81 ± 1.09 | 21.32 ± 0.69 |
TN(g100 g−1 cheese) | 5.15 ± 0.17 | 3.34 ± 0.11 |
WSN (% TN) | 10.34 ± 1.06 | 32.76 ± 0.95 |
TCA-N (% TN) | 6.37 ± 0.70 | 4.60 ± 0.04 |
TFAA(mgLeu/g) | 3.18 ± 0.56 | 3.72 ± 0.13 |
Physical-chemical parameters of mixed goat/ewes milk cheese from different geographical locations.
Adapted from Sulejmani et al. [31]
Most brine cheeses are dry-salted and are ripened and stored in brine, and the salting method is the basic difference in terms of varieties of cheeses. Traditionally, they are produced from sheep, cow, goat, or mixed raw milk [29, 30].
At the beginning the ripening of white-brined goat cheese, as1-CN (f24–199) and c2-casein were produced, indicating high activity of chymosin and plasmin. However, it can be seen that b-casein reduction rate was smaller than as1-casein that of during ripening (Figure 5). After 60 days of ripening in the GR cheeses, the band corresponding to as1-I-casein (as1-CN f102–191) was present in all electrophoretograms of the samples, as a result of hydrolysis of as1- casein. A reduction of as1- and b-casein was obviously faster in the GR cheeses than in the GP cheeses, probably due to the native microorganisms and indigenous milk enzymes. Significantly inactivated indigenous and milk proteinases indicates on great impact that had pasteurization [30].
(Left) Urea-PAGE of the water-insoluble fractions of white goat milk cheeses made using raw (GR) or pasteurized (GP) method during 120 days of ripening (right) (with permission from John Wiley and Sons) [10].
As it is obviously shown in Figure 5, the hydrolysis of as1-casein was faster in the GR cheeses during ripening obviously as a result of the higher activity of indigenous proteinases in the curd, which is exactly associated with the heat degree of the milk heat.
Some differences were observed during ripening for the fractions of peptides, which were eluted in the GR cheeses at higher quantity at the end of ripening than the beginning of ripening. Common peaks were evident in the 30 and 60 days of ripening in all of the chromatograms, with an increase in concentration of peptides during ripening, which were mainly eluted between the 56th and 76th min. In the chromatogram, between 64 and 74 min, the peak heights in the cheeses were generally much higher than in other cheese samples until the 120th day of ripening (Figure 6).
Reverse-phase HPLC profiles of the water-soluble fraction of white goat milk cheeses made using raw milk during 120 days of ripening [10].
The analysis of free amino acids in white-brined goat cheese confirmed the presence of all amino acids except tyrosine (Table 4). The quantity of free amino acids is low because of particular process of fermentation. Due to the high concentration of salt and low ripening temperature of white brine, the participation of thermophilic lactic acid bacteria in ripening is minimal, and this cheese is defined by a weaker breakdown of paracasein.
Free amino acids mg % | Ripening | |
---|---|---|
Day 15 | Day 60 | |
Lysine | 16.2 ± 0.31 | 33.11 ± 0.41 |
Histidine | 2.2 ± 0.13 | 16.12 ± 0.71 |
Arginin | 2.7 ± 0.71 | 4.85 ± 0.27 |
Threonine | 2.60 ± 0.78 | 3.71 ± 0.62 |
Valin | 11.78 ± 0.61 | 14.80 ± 0.16 |
Metionin | 6.11 ± 0.43 | 8.71 ± 0.62 |
Isoleucin | 3.21 ± 0.62 | 4.52 ± 0.27 |
Leucine | 11.72 ± 0.31 | 21.57 ± 0.36 |
Phenylalanine | 4.11 ± 0.62 | 15.61 ± 0.31 |
Total essential amino acids | 60.63 ± 0.64 | 123.01 ± 0.74 |
Asparagin acid | 3.8 ± 0.11 | 9.11 ± 0.37 |
Serin | 6.10 ± 0.18 | 12.11 ± 0.38 |
Glutamic acid | 6.70 ± 0.79 | 9.28 ± 0.65 |
Proline | 1.50 ± 0.28 | 3.61 ± 0.33 |
Glycine | — | 6.63 ± 0.58 |
Alanin | 3.31 ± 0.41 | 4.11 ± 0.43 |
Cistin | 4.81 ± 0.11 | 7.51 ± 0.38 |
Tirozin | — | — |
Total nonessential amino acid | 26.22 ± 0.53 | 52.36 ± 0.59 |
Total quantity | 89.18 ± 0.58 | 176.32 ± 0.71 |
The volatile components of white-brined goat cheeses have not previously been studied. They consisted of 12 acids, 14 esters, 6 ketones, 3 alcohols, 4 terpenes, and 6 miscellaneous compounds (Table 2). Acids, alcohols, and ketones constituted the principal chemical groups during ripening (mean volatile concentration of 51, 16, and 12% w/w of total compounds, respectively). The raw goat milk (GR) cheeses were by a higher quantity (78%) of total volatile compounds than the pasteurized goat milk (GP) cheeses, during ripening. Compared with day 1, a significant decrease in the total quantity of volatile compounds (except ketones and alcohols) was found after 120 days of ripening. Carboxylic acids are the principal volatile class in Macedonian white goat cheese (with 51% of total volatile compounds). The raw goat milk (GR) cheeses were characterized by higher quantity (86%) of total acids than the pasteurized goat milk (GP) cheeses during ripening. Milk heat treatment significantly (P < 0.05) influenced the concentrations of two volatile fatty acids (hexanoic acid and octanoic acid) (Table 2). Similarly, these compounds have been shown to be the principal volatile class in other goat milk cheeses [32]. The main acid was hexanoic acid (40% of total acids), and this was identified at significantly higher quantity in GR cheeses at 60 days of ripening. 3-Methylbutanoic acid was the most abundant branched chain fatty acid found in Macedonian goat cheese. This is in agreement with the findings of Beuvier et al. [33]. During ripening, total acids were at a higher concentration in the GR cheeses (86%) in comparison with GP cheeses (14%).
Caproic acid is a product of lipolysis, which significantly contributes to the smell of goat cheese [33]. Karagul et al. [34] explored the level of proteolysis in “Ezine” cheese, produced from a mixture of goat milk(40%), sheep’s milk (45–55%), and cow’s milk (up to 15%) without starter culture, during 8 months ripening. Urea-PAGE assay confirmed that αs-casein decomposes very quickly, while the β-casein degradations are almost constant. Differences in the rate of degradation are associated with pH and salt content [35].
Flavor is the main properties that influence the selection and consumption of cheeses. The effect of fatty acids on the sensory properties of different types of goat milk cheese is essential. The concentration of butyric acid is increased during the ripening of cheeses mainly higher of 50% from the total concentration in the beginning of its production (mozarella 66.35%, white-brined 74.58%, and pecorino 51.28%) [9]. The reason for the lower degree of formation of butyric acid during ripening and especially at the end of ripening of the cheeses is assumed to be the lack of a free substrate for conversion into fatty acids by way of lipolysis or reduction of enzyme activity due to the change in the microstructure of the cheeses.
At the first day of ripening in the raw goat milk cheeses (GR), acetic acid was identified at a higher concentration. Alcohols are the second most significant volatiles (16%) isolated in Macedonian goat cheese. At the end of ripening, the concentration of alcohols decreased to 40%, while after 60 days of ripening their quantity was 50%. The total esters were at higher quantity (98%) in raw goat milk (GR) cheeses than pasteurized goat milk (GP) cheeses during ripening. At the 60th day of ripening, a very high quantity of 3-methyl-1-butanol was found in the GR cheeses (Table 5). Heat treatment of the curd did not affect branched alcohols (except 2-ethyl-1-hexanol). 2-Propanone and 2-heptanone were the most abundant ketones among total of six ketones representing 54% and 13% of the total quantity of ketones, respectively. Higher quantity (77%) of total esters was characterized in the raw goat milk (GR) cheeses rather than the pasteurized goat milk (GP) cheeses during ripening. Because of particular odors and low perception thresholds, ester is very significant compounds in dairy products [36]. 2-Propanone and 2-heptanone were predominant ketones among six ketones that were identified in the Macedonian white goat cheeses.
Compounds | RI | Day 1 | Day 60 | P (type) | ||
---|---|---|---|---|---|---|
GP | GR | GP | GR | Type | ||
Acids | ||||||
2-Hydroxypropanoic acid | 8186 | 0.28 ± 0.40 | 101.04 ± 12.89 | ND | ND | NS |
2-Ethylbutanoic acid | 20,383 | 59.36 ± 8.95 | ND | ND | ND | NS |
Isobutyric acid | 26,446 | ND | 13.75 ± 9.44 | ND | ND | NS |
Butanoic acid | 28,317 | ND | 79.65 ± 12.64 | 47.89 ± 6.72 | 18.27 ± 9.73 | NS |
Pentanoic acid | 28,405 | ND | 8.18 ± 1.57 | ND | ND | NS |
3-Methyl, butanoic acid | 29,400 | ND | 1.35 ± 1.1 | 32.01 ± 3.64 | 16.19 ± 9.84 | NS |
Hexanoic acid | 33,722 | ND | 182.58 ± 19.81 | 46.86 ± 6.91 | 260.64 ± 127.86 | * |
Acetic acid | 37,692 | ND | 5.28 ± 7.47 | ND | ND | NS |
Octanoic acid | 38,524 | ND | 112.40 ± 18.96 | ND | 180.47 ± 174.55 | * |
Isobutyric acid | 41,235 | ND | 11.13 ± 1.74 | ND | ND | NS |
Decanoic acid | 42,882 | ND | 45.36 ± 6.14 | ND | ND | NS |
2-Ethyl, caproic acid | 45,717 | ND | 17.62 ± 13.39 | ND | 10.68 ± 5.10 | NS |
Total | 59.6 ± 9.3 | 578.3 ± 47.9 | 126.7 ± 16.2 | 486.2 ± 324.1 | ||
Ketones | ||||||
2-Propanone | 6352 | 18.19 ± 3.11 | 64.82 ± 37.66 | 12.82 ± 5.71 | 51.55 ± 33.87 | * |
2-Butanone | 7706 | 1.68 ± 0.49 | 4.23 ± 1.21 | 3.89 ± 0.26 | 3.76 ± 1.24 | NS |
2-Pentanone | 9446 | 3.40 ± 2.46 | 7.22 ± 4.97 | 6.65 ± 1.36 | 8.65 ± 3.60 | * |
2-Heptanone | 15,472 | 1.93 ± 2.16 | 3.48 ± 3.12 | ND | 19.87 ± 7.07 | NS |
2-Octanone | 15,501 | ND | ND | 0.98 ± 0.39 | ND | NS |
2-Nonanone | 21,938 | ND | ND | ND | 8.38 ± 7.24 | NS |
Total | 25.12 ± 12.3 | 79.15 ± 55.8 | 24.3 ± 13.3 | 92.5 ± 55.9 | ||
Esters | ||||||
Methyl acetate | 6521 | 3.35 ± 1.90 | 9.12 ± 1.90 | 3.17 ± 1.58 | 6.51 ± 1.12 | * |
Ethyl acetate | 7447 | 19.43 ± 14.40 | 2.14 ± 3.03 | 2.24 ± 1.88 | 6.18 ± 6.79 | NS |
Methyl propanoate | 7823 | 1.86 ± 0.86 | 4.56 ± 0.58 | 2.30 ± 1.69 | 3.71 ± 0.21 | NS |
Methyl butyrate | 9645 | 2.28 ± 0.69 | 12.98 ± 5.42 | 3.69 ± 4.12 | 14.19 ± 6.39 | * |
Methyl carbonate | 9785 | 0.15 ± 0.21 | 2.70 ± 0.18 | ND | ND | NS |
Ethyl butyrate | 11,035 | 1.65 ± 2.34 | ND | ND | ND | NS |
n-Butyl acetate | 12,024 | 1.04 ± 1.47 | ND | ND | ND | NS |
Isoamyl acetate | 13,535 | 1.59 ± 2.24 | ND | ND | ND | NS |
Methyl caproate | 15,544 | 0.34 ± 0.49 | 2.56 ± 3.63 | ND | ND | NS |
Ethyl heptanoate | 15,973 | ND | 23.29 ± 32.93 | ND | ND | NS |
Isoamyl acetoacetate | 16,141 | 0.72 ± 1.02 | ND | ND | ND | NS |
Ethyl n-caproate | 17,021 | 0.17 ± 0.25 | ND | ND | ND | NS |
Dimethyl phthalate | 24,779 | 1.74 ± 2.46 | 3.16 ± 4.47 | ND | ND | NS |
Diethyl phthalate | 45,468 | 1.77 ± 1.18 | 25.06 ± 26.41 | 1.25 ± 1.77 | 1.11 ± 1.56 | NS |
Total | 36.10 ± 29.51 | 85.58 ± 78.56 | 12.65 ± 11.04 | 31.70 ± 16.07 | ||
Terpenes | ||||||
dl-Limonene | 15,942 | 54.96 ± 7.07 | ND | 109.79 ± 31.99 | ND | * |
Cymene <para-> | 18,287 | 1.80 ± 2.54 | ND | ND | ND | NS |
Alph.-thujene | 14,771 | 0.10 ± 0.14 | ND | ND | ND | NS |
Alpha-pinene | 10,644 | 15.61 ± 10.93 | 14.80 ± 6.62 | 49.14 ± 30.44 | 6.66 ± 0.37 | NS |
Total | 72.47 ± 20.67 | 14.80 ± 6.62 | 158.92 ± 62.44 | 6.66 ± 0.37 | ||
Alcohols | ||||||
Ethanol | 8302 | 1.79 ± 0.05 | 3.85 ± 5.44 | ND | 1.70 ± 1.10 | NS |
3-methyl, 1-butanol | 16,117 | 1.06 ± 1.50 | 49.01 ± 5.25 | ND | 233.12 ± 155.11 | * |
1-Pentanol | 17,429 | ND | 2.66 ± 0.24 | ND | ND | NS |
Total | 2.84 ± 1.54 | 55.52 ± 11.93 | ND | 234.81 ± 156.22 | ||
Miscellaneous | ||||||
Pentane | 4767 | 0.24 ± 0.34 | 6.08 ± 8.60 | 5.49 ± 4.21 | ND | NS |
Hexane | 4905 | 7.67 ± 3.73 | 2.33 ± 0.31 | 4.56 ± 1.43 | 3.34 ± 2.89 | NS |
Dimethyl sulfide | 5742 | ND | 6.56 ± 9.28 | 3.44 ± 3.20 | ND | NS |
Methylamine-D2 | 7583 | 1.22 ± 1.72 | 6.76 ± 0.88 | 9.96 ± 4.63 | 8.05 ± 4.23 | NS |
2-Methylbutanal | 7976 | 0.48 ± 0.43 | 4.90 ± 4.14 | ND | 2.96 ± 3.21 | * |
3-Methylbutanal | 8064 | 4.10 ± 3.57 | 35.28 ± 12.13 | 3.94 ± 1.56 | 29.09 ± 30.27 | ** |
Total | 13.7 ± 5.7 | 61.5 ± 26.4 | 27.3 ± 9.3 | 45.0 ± 37.7 | NS |
Mean values ± SD of volatile compounds identified in pasteurized (GP) and raw (GR) white goat’s milk cheeses after 1 and 60 days of ripening (μg/100 g).
P < 0.05.
P < 0.01.
Mean data for three batches of pasteurized (GP) and raw (GR) goat cheese analyzed in triplicate. RI, retention index; ND, not identified; NS, not significant; P, probability. P age is probability for ripening period (i.e., 1, 60, or 120 days), P type is probability for cheese type (i.e., GP or GR) [10].
Adapted from Sulejmani and Hayaloglu, 2017
Five different acids that were identified in the Goat beaten cheese from Kumanovo region with concentration of 13347.2 μg 100 g−1 were reported. Six different acids were identified in the goat beaten cheese from the Radoviš region, and their concentration were 13773.8 μg 100 g−1, respectively [31]. Hexanoic, octanoic, and decanoic acids were responsible for the characteristic aroma of goat cheeses, and their contribution to the volatile profile of beaten cheeses has been shown in this study as well, giving rise to the trivial terms caproic, caprylic, and capric acids, respectively. In addition 2-heptanone was identified at highest concentration in Kumanovo beaten cheese than other regions (Figure 7). Also 2-octanone, 8-nonen-2-one, and 2-nonanone were rarely found in cheeses from other regions.
GC-MS chromatogram of volatiles compounds identified form goat beaten cheese [21].
Heptanol is determined in highest amount in the beaten cheese with goat milk from the northeast region and has been identified as a key aromatic flavor- component in gorgonzola and grana padano cheese [37]. Curioni and Bosset [38] reported a high concetration of heptanols as well as in semihard varieties of Spanish cheese from goat’s milk.
The breeding of the goats in the Republic of Macedonia has a mark of tradition, namely, because of appropriate grazing conditions. This is very economical, because the entire diet is reduced to a grazing with less extra feed in the form of concentrate and minerals. Also Macedonia has real possibilities for transformation of many extensive goat farms into an organic farm. In addition, it creates additional opportunities for the outlay on the markets in the EU, for which the organic production year-by-year increases. Macedonian goat cheeses are being differentiated by their strong bounds with the territory of their origin, and so they represent a historical and cultural designation of the community which they are produced by. Production of these cheeses is in limited geographical areas with use of know-how techniques transferred from generation to generation and use of milk that has undergone no treatment after milking. Goat milk can be successfully used to produce various cheeses because cheese is characterized by a specific lactic acidic taste and aroma, a good degree of protein breakdown. Lactic acid processes and changes in moisture content take place more evenly during ripening than cow cheeses. From milking to the end of ripening process, this type of cheeses is passing through different surroundings where a variety of microorganisms have an opportunity to grow and develop. The research that has been conducted has shown that traditional made cheeses have unique benefits in terms of palatable pleasure, richness, and diversity as well as protection against pathogens. Undoubtedly their properties have been achieved due to the presence of unique indigenous microbiota especially because of the use of raw milk, combined with specific skills that give their general characteristic properties and quality. In order to understand the situation of traditional milk processing and utilization in this part of the state, one should recall that milk production has an obvious seasonality related to climatic conditions and most of these products are homemade following neither standardized conditions nor proper hygiene standards. Careful attention must be paid to hygiene in order to produce milk of high bacteriological quality. However, despite all precautions, it is impossible to completely exclude bacteria from milk. Therefore, good hygiene is particularly important in producing especially fresh ripened cheeses. So prevention of contamination of the milk and meticulous attention to good hygiene during cheese production and ripening will reduce the incidence of pathogens; therefore, good acid-produced cheese during proper ripening is also helpful.
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