Sea salts as a potential source of food spoilage fungi

Food Microbiology 69 (2018) 89e95

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Food Microbiology journal homepage: www.elsevier.com/locate/fm

Sea salts as a potential source of food spoilage fungi Megan N. Biango-Daniels*, Kathie T. Hodge Plant Pathology and Plant-Microbe Biology, School of Integrative Plant Science, College of Agriculture & Life Sciences, Cornell University, 334 Plant Science Building, 236 Tower Road, Ithaca, NY 14853-5904, USA

a r t i c l e i n f o

a b s t r a c t

Article history: Received 31 March 2017 Received in revised form 26 June 2017 Accepted 31 July 2017 Available online 1 August 2017

Production of sea salt begins with evaporation of sea water in shallow pools called salterns, and ends with the harvest and packing of salts. This process provides many opportunities for fungal contamination. This study aimed to determine whether finished salts contain viable fungi that have the potential to cause spoilage when sea salt is used as a food ingredient by isolating fungi on a medium that simulated salted food with a lowered water activity (0.95 aw). The viable filamentous fungi from seven commercial salts were quantified and identified by DNA sequencing, and the fungal communities in different salts were compared. Every sea salt tested contained viable fungi, in concentrations ranging from 0.07 to 1.71 colony-forming units per gram of salt. In total, 85 fungi were isolated representing seven genera. One or more species of the most abundant genera, Aspergillus, Cladosporium, and Penicillium was found in every salt. Many species found in this study have been previously isolated from low water activity environments, including salterns and foods. We conclude that sea salts contain many fungi that have potential to cause food spoilage as well as some that may be mycotoxigenic. © 2017 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

Keywords: Sea salt Fungal spoilage Ingredient safety Filamentous fungi

1. Introduction Valued for their nuanced taste and texture and their often artisanal origins, sea salts have lately seen a surge in popularity (Parks, 2014). Salt is widely perceived to be a chemically pure and sterile food ingredient, however, sea salts may carry microbial contaminants (Butinar et al., 2011). Sea salt was shown to be the source of a mycotoxin-producing mold that spoiled dry-cured meat in a Slovenian production facility (Sonjak et al., 2011a). This finding, and the ways in which sea salts are produced and handled, raised the question of the presence of spoilage fungi in sea salts. One reason sea salts may contain viable fungi is the timehonored method by which salt is harvested from seawater by slow evaporation in shallow ponds called salterns. A third of all salt for human consumption is produced this way (Butinar et al., 2011). Some organisms in saltern ecosystems may persist as viable propagules even when water activity becomes prohibitive to growth. Thus, sea salt may be enriched in microbes that can grow in high salt environments e the very conditions relied on as an important principle of food preservation (Butinar et al., 2011;

* Corresponding author. E-mail addresses: [email protected] (M.N. Biango-Daniels), KH11@cornell. edu (K.T. Hodge).

Gostincar et al., 2010; Grant, 2004; Gunde-Cimerman et al., 2009, 2000; Gunde-Cimerman and Zalar, 2014a; Javor, 2002; Jay et al., 2008; Zajc et al., 2014). This study aimed to quantify viable filamentous fungi in sea salts that could cause spoilage when sea salts are used as food ingredients. A selective medium was used to capture living, fastgrowing, moderately xerophilic fungi in commercial sea salts. The fungi were identified by DNA sequencing, and fungal communities were examined relative to their ocean of origin. 2. Materials and methods 2.1. Sampling This study aimed to assess the fast growing, moderately xerophilic filamentous fungi present in sea salt, which are likely to spoil a range of salted products. Seven salts were sampled, representing a variety of commercially available “Sea Salts” in the United States. Duplicate packages were purchased of three Atlantic Ocean salts from France (Fr1, Fr2, and Fr3), three Pacific Ocean salts (Us1, Us2 and Haw), and one mined salt from the Himalayan Mountains (Him). While six of the salts were made from ocean waters, the seventh (Him) was labeled as a sea salt when in fact it was mined from an ancient sea salt deposit: this distinction is noted throughout our analysis. Each duplicate package (true replicates)

http://dx.doi.org/10.1016/j.fm.2017.07.020 0740-0020/© 2017 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

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was sampled two or three separate times, for a total of five subsamples (pseudoreplicates) per type of salt. The salts varied in their appearance and textures. The Atlantic salts (Fr1, Fr2, and Fr3) were fine-grained white or grey salts with no additives. Two of the Pacific salts (Us1 and Us2) were white, fine-grained finishing salts with the additives calcium silicate, dextrose, potassium iodide in both and sodium bicarbonate in one (Us2). The third Pacific salt (Haw) was a black, medium-coarse salt with no additives. The one mined salt (Him) included was a very coarse, dark pink salt. 2.2. Isolation Fungi were isolated from each of the 35 subsamples of sea salts (five per each of the seven types of salt). Each subsample was weighed and dissolved in 100 ml of room temperature sterile deionized water and immediately filtered through disposable MicroFunnel (Pall) filter funnels with Supor membranes (0.2 mm pore size), all under aseptic conditions. Sterile deionized water without added salt was used as a negative control. The filtered particles, including fungi, were trapped in the membranes, which were removed and placed filtrate-side-up on dichloran-glycerol (DG18) enumeration medium supplemented with 30 mg ml 1 streptomycin sulfate (Butinar et al., 2011; Gunde-Cimerman et al., 2009, 2003, 2000; Hocking and Pitt, 1980). This medium was developed to isolate moderately xerophilic food spoilage fungi. Plates were incubated at room temperature, 24 C (±2 ), for one week. Although the short incubation did not allow for the detection of slow-growing xerophiles, it was ideal to detect fast-growing fungi more likely to induce spoilage concerns. Fungal colonies that emerged in seven days were counted; all were removed aseptically and subcultured on DG18 to obtain unique individual isolates, determined by colony morphology and microscopic characteristics. 2.3. DNA extraction and PCR amplifications Mycelia harvested from DG18 plates were physically disrupted using a pestle in a 1.5 mL microcentrifuge tube containing sterile 0.5 mm glass beads (Biospec Products). Genomic DNA was extracted using the PrepMan Ultra Sample Preparation Reagent kit (Applied Biosystems by Life Technologies), E.N.Z.A. SP Fungal DNA Mini Kit (Omega Bio-Tek), or DNeasy Plant Mini Kit (Qiagen) following recommended protocols. Extracted genomic DNA was stored in elution buffer at 20 C. DNA from individual isolates was the target of PCR amplification of the internal transcribed spacer (ITS) regions of the rDNA repeat, a widely accepted barcode for fungi (Schoch et al., 2012). The forward ITS5 primer (50 -GGAGTAAAAGTCGTAACAAGG -30 ) and reverse ITS4 primer (50 -TCCTCCGCTTATTGATATGC-30 ) (White et al., 1990) were used in 25 mL reactions with Quanta AccuStat Taq DNA polymerase or Sigma-Aldrich REDTaq genomic DNA polymerase with MgCl2. A DNA Engine PTC-200 thermal cycler (MJ Research) was programmed as follows for the REDTaq PCR: initial denaturation 95 C/ 2 min, 26 cycles of 94 C/1 min, 55 C/1min, and 72 C/3 min, and final extension of 72 C/15 min. For the AccuStat Taq PCR: initial denaturation 95 C/2 min, 20 cycles of 94 C/1 min, 55 C/30 s, and 72 C/1 min, 15 cycles of 94 C/30 s, 60 C/30 s, and 72 C/1 min, and final extension of 72 C/10 min. 2.4. Isolate identification PCR amplicons were cleaned up using E.N.Z.A. Cycle-Pure (Omega Bio-tek) according to manufacturer instructions and sequenced with a PCR primer using an Applied Biosystems Automated 3730xl DNA analyzer at the Cornell University Biotechnology

Resource Center. Isolate sequences were identified with NCBI nucleotide BLAST (Agostino, 2012; Altschul et al., 1990). Nucleotide sequences were first blasted (Megablast) against all fungal (taxid: 4751) sequences, excluding uncultured/environmental sample sequences. Because of uncertainty about reputability of the identities of some NCBI accessions, Aspergillus and Penicillium sequences were identified by blasting against a “gold-standard” ITS barcode database assembled from GenBank accession numbers selected by the International Commission on Penicillium and Aspergillus (Samson et al., 2014; Visagie et al., 2014b). Cladosporium isolate identities were inferred to the level of species complex by blasting against taxonomically validated sequences from a recent taxonomic monograph (Bensch et al., 2012). Isolate identities were inferred from among BLAST results with a zero E (expect) value, indicating a significant match, by selecting the match with the highest max score, which accounts for quality and length of the alignment (Agostino, 2012). Isolates that matched multiple species equally well were analyzed with the MOLE-BLAST neighbor-joining treebuilding tool, a multiple alignment program based on MUSCLE (Edgar, 2004; Saitou and Nei, 1987).

2.5. Statistical analysis To determine whether the quantity of fungal isolates in a sample was associated with salt type, a generalized linear mixed model (GLMM), fitted with a Poisson model, which is appropriate for discrete, positive responses such as counts, was used (O'Hara, 2009). The GLMM included a fixed effect of salt type and random effect of salt package, as there were multiple subsamples from each of two packages of every salt type. Weight was included as an offset in the model because subsample weights were not identical, ranging from 2.9 to 5.4 g with a mean weight of 3.27 g, and more isolates would be expected from larger subsamples. The data were found to be overdispersed (greater observed variance than expected by the model), so a random effect at the observation level , 2015). The effect of salt type on was included (Lenth and Herve quantity of fungi was assessed with a likelihood ratio test, which was compared to an intercept-only model. These models were executed using the lme4 package in R statistical programming software (Bates et al., 2014; R Core Team, 2015). A post-hoc pairwise comparison of mean CFU/g fungi among the seven salts was performed. Tukey's adjustment was also performed to avoid false , positives caused by multiple comparison issues (Lenth and Herve 2015). To compare the fungal community composition among the seven salts and between salts from Atlantic and Pacific oceans, excluding the mined salt, ordination was done on the six species that were found in more than one subsample. Ordination is a dimension reduction technique that allows visualization of salt communities' similarities by their proximity to each other in a twodimensional ordination space (Ramette, 2007). Here, nonmetric multidimensional scaling (NMDS) was used with the Bray-Curtis dissimilarity index (Ramette, 2007). Species isolated from only one of the salts sampled (including all the Penicillium species) were not included, as they cannot contribute to assessments of similarities among subsamples. Subsamples that contained no species or contained only unique species were excluded, leaving 16 subsamples that were analyzed. Before analysis the data were relativized by subsample total so that each species' value was its percent abundance in the subsample. Ordination was executed using the Vegan package in R (Oksanen et al., 2015). The ANOSIM function was used to test for significant differences in communities of different salt types (Ramette, 2007).


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3. Results

3.2. Fungi in sea salts

3.1. Quantification of viable fungi in sea salts

Species composition varied among sea salts but salts from different oceans were united by shared species mainly in the genera Aspergillus and Cladosporium (Fig. 2). The most abundant genera were Aspergillus, Cladosporium, and Penicillium, which each contributed multiple species. No Penicillium species was common among two or more salts (Fig. 2). Species composition varied among salts but several Aspergillus and Cladosporium species were found in multiple salts. In contrast to those shared taxa, 13 taxa were found in low abundance or in only one salt (Fig. 2). Sixteen fungal species and three different Cladosporium taxa, representing seven genera, were found among the seven salts (Table 1). Mined salt, Him, had no fungi in common with any of the sea salts and yielded only one isolate of Penicillium camemberti (Fig. 2). The six taxa found in multiple sea salts were: Aspergillus fumigatus, A. niger, A. sydowii, and A. welwitschiae, and the Cladosporium cladosporiodes and C. herbarum complexes (Fig. 2). Ordination analysis of taxa from different salts found no significant differences among fungal communities (ANOSIM R ¼ 0.066, p ¼ 0.24).

Of the thirty-five subsamples of sea salts sampled, thirty-two of them contained viable fungi, and there were fungi present in each sea salt. A total of eighty-five fungal isolates were obtained among the subsamples in this study (Table 1). The mean number of viable fungal propagules in each salt, measured in colony forming units per gram (CFU/g), ranged from 0.07 to 1.71 (Fig. 1). Looking at the ocean of origin for the salts, there was no clear difference in number of viable fungi (Fig. 1). The salts with the fewest propagules were the mined salt, Him, and Us1, with 0.066 and 0.126 CFU/g respectively, while the salt with the greatest CFU/g was Fr2 (Fig. 1). There was significant variation among salt types in number of fungi (Fig. 1), which did not show a package effect (likelihood ratio test X26 ¼ 14.875, p ¼ 0.021). The total number of isolates from the combined subsamples was highest in Fr2, which yielded 26 isolates (Fig. 2).

Table 1 Viable fungi isolated from seven commercially available salts and corresponding NCBI accession number. The isolates were deposited in the Food Fungi Collection at the New York State Agricultural Experiment Station as frozen cultures. Ocean

Salt

Isolate number

Best hit (BLAST match)

NCBI Accession

Atlantic

Fr1

1100 1052 1030 1031 1060, 1065 1077 1073 1043 1039A, 1040, 1049, 1054A, 1097

Aspergillus chevalieri (EF652068) Aspergillus fumigatus (EF669931) Aspergillus sydowii (EF652450) Aspergillus xerophilus (EF652085) Cladosporium cladosporioides complex (FJ936159) Cladosporium cladosporioides complex (HM148003) Penicillium cremeogriseum (GU981586) Aspergillus niger (EF661186) Aspergillus sydowii (EF652450)

1017, 1025, 1026, 1039B, 1041, 1042, 1044, 1054B

Aspergillus welwitschiae (FJ629340)

1038 1058, 1059, 1062A, 1066, 1074, 1083

Aspergillus unguis (EF652443) Cladosporium cladosporioides complex (FJ936159)

1088 1062B, 1067, 1068 1024 1029 1069 1018, 1019, 1027, 1028, 1045, 1046, 1047, 1048, 1050, 1051, 1052

Cladosporium cladosporioides complex (HM147999) Cladosporium herbarum complex (EF679334) Ulocladium dauci (NR_077186) Aspergillus niger (EF661186) Aspergillus tubingensis (EF661193) Aspergillus welwitschiae (FJ629340)

1061 1071, 1079 1055 1075 1015, 1016, 1032 1033A, 1056A 1056B 1072, 1080 1057, 1064, 1070 1023, 1033B, 1034, 1035, 1036, 1037

Cladosporium herbarum complex (EF679334) Aspergillus caespitosus (EF652428) Aspergillus fumigatus (EF669931) Calostilbe striispora (KM231855) Cladosporium cladosporioides complex (HM148003) Cladosporium cladosporioides complex (FJ936159) Cladosporium cladosporioides complex (HM147994) Cladosporium cladosporioides complex (FJ936159) Cladosporium sphaerospermum complex (DQ780364) Penicillium oxalicum (AF033438)

1076 1095, 1084, 1089 1092 1094 1081, 1090 1098, 1086

Scopulariopsis sphaerospora (LM652438) Aspergillus welwitschiae (FJ629340) Cladosporium cladosporioides complex (FJ936159) Cladosporium cladosporioides complex (HM147994) Cladosporium cladosporioides complex (HM147999) Cladosporium cladosporioides complex (HM147998) Cladosporium herbarum complex (EF679334) Cladosporium herbarum complex (JN906977) Septoriella phragmitis (KR873251) Penicillium camemberti (AB479314)

KT826690 KT826642 KT826618 KT826619 KT826651, KT826669 KT826665 KT826633 KT826628, KT826643, KT826608, KT826629, KT826634, KT826627 KT826649, KT826658, KT826678 KT826654, KT826612 KT826617 KT826661 KT826609, KT826616, KT826637, KT826641, KT826652 KT826663, KT826645 KT826667 KT826606, KT826621, KT826647 KT826664 KT826648, KT826611, KT826624, KT826668 KT826685, KT826675, KT826682 KT826682 KT826684 KT826672, KT826680 KT826688, KT826676

Atlantic

Atlantic

Pacific

Pacific Pacific

Mined

Fr2

Fr3

Haw

Us1 Us2

Him

1096 1091, 1093

1082, 1087 1099

KT826652

KT826630, KT826639, KT826687 KT826613, KT826614, KT826631, KT826632, KT826644 KT826650, KT826653, KT826666, KT826674 KT826659, KT826660

KT826610, KT826615, KT826635, KT826636, KT826638, KT826640, KT826642 KT826670

KT826607, KT826620 KT826646

KT826656, KT826662 KT826622, KT826623, KT826625, KT826626 KT826686 KT826681, KT826683

KT826673, KT826677 KT826689

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Fig. 1. Mean viable fungal propagules (C.F.U./g) with standard error for each of the seven salts, organized by ocean of origin. Salts that have the same letter are not significantly different from each other (a ¼ 0.05) when no corrections were made for multiple hypotheses tests. After using Tukey's method for correction, only Fr2 and Us1 were significantly different (a ¼ 0.05).

Similarly, a broader analysis based on ocean of origin also found no significant differences (ANOSIM R ¼ 0.012, p ¼ 0.46) (Fig. 3).

long periods, posing a potential downstream food spoilage risk if they can grow in moderately low water activities common for most foods.

4. Discussion 4.2. A global saltern community of fungi 4.1. Viable fungi in sea salts Commercial sea salts consistently contained low levels of a variety of living fungi. This study probably underestimated the true abundance of fungi in sea salt, since only one growth medium was used and very slow-growing fungi were excluded by the one-week incubation time. Fungi were identified using ITS barcoding, which generally works well, although for a few groups species complexes are difficult to resolve (Schoch et al., 2012). Some of the detected fungi have the potential to cause food spoilage when sea salt is used as a food ingredient, especially in products that do not receive heat treatment, as is the case in dry-cured meats in which sea salts were shown to be the source of spoilage mold inoculum (Sonjak et al., 2011a). The origins of these sea salt fungi are uncleard they might arise during production directly from the saltern environment, or by introduction of fungi during processing, handling, or packing. The later source is likely where the single fungal isolates in the mined salt originated. Sea salts are produced from seawater trapped in solar salterns: flow-through networks of exposed, shallow ponds (Javor, 2002). As sea water is progressively concentrated through evaporation, the increasingly saline waters harbor distinctive communities of microbes including bacteria, archaea, algae and fungi (Food and Drug Administration, 2012; Javor, 2002). These xerophilic species of endemic saltern communities can be carried over into finished salt (Butinar et al., 2005), where they may even contribute to the distinctive flavor, texture, and terroir of sea salts. Fungal propagules that can survive contact with salt may persist in the product for

Statistical analysis supports the idea of a pan-global saltern community, in that there was no evidence that sea salt communities are distinct to the Atlantic vs. Pacific oceans (Fig. 3). Even salts from very distant locations were not significantly different in their fungal composition. Species of Penicillium, Cladosporium, and Aspergillus predominate in the sea salts examined, and also in salterns (Gunde-Cimerman and Zalar, 2014b). The tested salts contained aspergilli already reported from global saltern communities, including A. niger, A. sydowii, A. chevalieri, A. tubingensis, and A. fumigatus (Fig. 2) (Butinar et al., 2011). Aspergillus nidulans and A. chevalieri are known in other low water activity environments, including foods and salterns (Gesheva and Negoita, 2011; Vytrasova et al., 2002). Most aspergilli found in salt belong to Aspergillus section Nigri, a group of dark-spored species known for their global saltern distribution and ability to spoil low water activity foods (Butinar et al., 2011; Pitt and Hocking, 2009). Due to their cosmopolitan nature it is impossible to determine whether all the detected isolates originated in salterns or were airborne contaminants (Butinar et al., 2005; Visagie et al., 2014a). Aspergillus welwitschiae was abundant in several salts and may be a previously unidentified saltern fungus. This is the first report of Cladosporium species from finished sea salt. Cladosporium is a prolific, cosmopolitan genus associated with solar salterns and other low water activity environments, as well as indoor and outdoor air (Butinar et al., 2005; Cantrell et al., 2013; Gunde-Cimerman et al., 2003, 2000; Schubert et al., 2007; Zalar et al., 2007).


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Fig. 2. Total number of isolates from the combined subsamples of each salt ranged from 1 in mined Him salt to 26 in Fr2. Fungi that were found in only one salt are indicated by * in the legend. Colors correspond to taxa found in each salt, displayed left to right by ocean of origin, with the mined salt, Him, on the far right.

Fig. 3. Ordination analysis of the global distribution of fungi across salt subsamples grouped by ocean of origin, excluding mined Him salt. Overlapping ellipses, generated by the R package Vegan, represent the standard deviation of NMDS coordinates of samples grouped by ocean of origin and illustrate that fungal community compositions between Atlantic and Pacific salts were not significantly different (p ¼ 0.46).

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Based on their plant pathogenic ecology, Calostilbe striispora and Septoriella phragmitis were likely introduced from plant populations adjacent to salterns or salt storage facilities (Crous et al., 2015; Rossman et al., 1999). Saprobes such as Ulocladium dauci and Aspergillus caespitosus, known from soil and decay communities, were likely introduced from soil contact during harvest (Runa et al., 2009; Samson and Mouchacca, 1975). Many penicillia, including P. oxalicum, are recognized as locally abundant saltern species that dominate their niche (Butinar et al., 2011) (Fig. 2). Penicillium cremeogriseum is unknown from salterns but recognized as an indoor air contaminant (Ejdys and Biedunkiewicz, 2011). It may have been introduced during salt storage after harvest. Him salt, the only mined type of salt in this study, yielded only one isolate which must have been introduced during processing or storage (Fig. 2).

4.3. Salt fungi as spoilage risks Fungi detected in sea salts in this study have the significant potential to spoil foods, and introduce mycotoxins or allergens when sea salt is used as a food ingredient (Bennett and Klich, 2003; Butinar et al., 2011; Frisvad et al., 2004; Ponsone et al., 2007; Steyn, 1969). Penicillium oxalicum and Scopulariopsis sphaerospora, for example, are important spoilers of sausage and other low water activity foods, and may be introduced to food through salt (Canel et al., 2013; Iacumin et al., 2009; Papagianni et al., 2007; Wolfe et al., 2014). Penicillium camemberti was isolated from the mined salt and is used to ripen cheeses, but some strains that may be introduced via sea salt produce mycotoxins (Frisvad et al., 2004; Ropars et al., 2012) and so are undesirable in cheese production. Some Cladosporium spp. cause meat spoilage problems in factories and might also originate from sea salts used as ingredients (Hammami et al., 2014; Mandeel, 2005; Sonjak et al., 2011b; Sørensen et al., 2008). Multiple species detected in this study, including A. nidulans, A. fumigatus, A. caespitosus, and A. niger, and P. €se et al., camemberti are capable of producing mycotoxins (Bra 2009). All sea salts examined in this small study contained living fungi. This is concerning given that there are no microbiological standards of quality for salt according to the Compendium of Methods for the Microbiological Examinations of Foods and the Codex Alimentarius of Food and Agricultural Organization (Downes and Ito, 2001; Alimentarius, 1985). Standards are needed, as are methods to reduce the risks of sea salts as spoilage mold sources, which might include simple modifications to production and handling practices, such as reducing soil contact and improving storage conditions. Future work will elucidate the full range of fungi in diverse sea salts, and investigate their origins and mitigation in sea salt production.

Acknowledgments We are grateful to Erika Mudrak of the Cornell Statistical Consulting Unit for her data analysis advice and expertise, to Katie Wilkins for her assistance creating databases used for identification of Aspergillus, Penicillium, and Cladosporium isolates. We thank Dr. Randy Worobo and Dr. Eric Nelson of Cornell University, and anonymous reviewers for their comments on this manuscript. This work was supported by the USDA National Institute of Food and Agriculture, Hatch project 1002546. Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the authors and do not necessarily reflect the view of the National Institute of Food and Agriculture (NIFA) or the United States Department of Agriculture (USDA).

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