The impact of freezing on the post-mortem human microbiome

Introduction

The use of human bodies in research dates as far back as the 16th century, with the dissection of cadavers for understanding the human anatomy (Varner et al., 2021). Consequently, human corpses have been integrated into teaching for training medical practitioners (Habicht et al., 2018; Holm and Iaizzo, 2019). The study of human cadavers has also been adopted in other disciplines, including forensics – for the purpose of developing and validating new methods to assist forensic experts in death investigations (Gunderman and Wilson, 2005; Shin and Meals, 2005; Turney, 2007; Anyanwu, 2010; Varlet et al., 2020).

Forensic taphonomy, which aims to understand human (or animal, when used as proxies) decomposition over time (Miles et al., 2020; Martin et al., 2021), is a key area of forensic research that more often has utilized human donors for essential research. The importance of such research has been cited in multiple publications (Iqbal et al., 2018; Miles et al., 2020), as this can provide answers to fundamental questions regarding post-mortem interval (PMI) or post-burial intervals, as well as the cause and/or circumstances of death (Miles et al., 2020). Incorporating human donors in forensic taphonomy research can be vital to reaching conclusions and informing standard practice for PMI estimation (Fancher et al., 2017; Matuszewski et al., 2020), discovery of clandestine depositions (Pringle et al., 2020; Doro et al., 2022) and training human remains detection dogs (Komar, 1999; Cablk et al., 2012). This is particularly crucial, as animal replacements may not fully represent true decomposition processes seen in humans (Williams et al., 2019b; Matuszewski et al., 2020). Thus, human taphonomy facilities – where human donors are studied for the purpose of forensic research (Williams et al., 2019a,b) – have been established in different countries for these purposes, among others, and have become centres for multidisciplinary research.

One of such areas of research involves the study of the microbial communities existing on and within human donors, and their related environments during decomposition. Microbiome investigations in forensic human taphonomy studies primarily focus on examining bacterial (and other microbial) succession during human decomposition for predicting PMI or cause of death. However, significant changes to microbial communities and their interaction networks can be driven by even small changes to temperature (Carter et al., 2008; Metcalf, 2019; Burman and Bengtsson-Palme, 2021), setting back our understanding of these succession patterns. Thus, recommendations for sampling in microbiome studies – forensic or otherwise – often suggest the immediate extraction of microbial DNA or the preservation of samples when immediate extraction is not possible (Lauber et al., 2010).

Sample preservation for stabilizing the microbiome often involves storing samples at below freezing temperatures, among other practices. Freezing at −80°C is often considered the gold standard for long-term preservation of microbiome-related samples, particularly fecal matter (Choo et al., 2015; Vandeputte et al., 2017). However, immediate preservation of samples at this temperature is not always feasible, especially at scenes of forensic interest. Hence, some studies have assessed the effect of freezing samples – soil (Bhattacharjee et al., 2021; Ouyang et al., 2021), fecal matter (Choo et al., 2015), saliva (Furuhashi et al., 2022) – at various temperatures and the influence on their associated microbiomes, with conflicting results. Even with such inconsistent findings, it is common practice to freeze these samples, frequently at −20°C, to limit microbial activity or proliferation.

Similarly at human taphonomy facilities, preserving donors by freezing prior to the commencement of decomposition research is relatively common practice, particularly in situations where multiple bodies must be deposited at the same time for decomposition studies involving more than one corpse. Thus, freezing is often used to halt or slow decomposition, and therefore maintain the bodies at a PMI hypothetically close to zero. In meat science studies, it has generally been determined that freezing and thawing do not effectively change the number of viable microbes present in meat (Vieira et al., 2009; Leygonie et al., 2012; da Silva Bernardo et al., 2020). However, it must be noted that these studies primarily assess microbial presence through culture studies. Yet there is information on negative effects of non-optimal freezing due to changes in osmotic pressure (Baust et al., 2009; Li et al., 2018). This can result in the rupture of cells through damaged cell walls which could release sequestered microbes. While the impact of freezing on the microbiome of forensic research samples has been explored, there is no information – to the best of our knowledge – on how freezing whole human donors may impact their microbiome.

Understanding any effects of freezing is vital for forensic human taphonomy research, as shifts to the microbial community composition and abundance prior to the start of a study may influence the results and therefore the interpretation of results. Previously, studies have aimed to understand decomposition related changes to microbial activity in cases where animal carcasses or sections of tissues have been frozen prior to burial. One of such studies reported that freezing plays a significant role in the type of decomposition observed – aerobic decomposition in previously frozen animal cadavers and anaerobic in fresh cadavers (Micozzi, 1986). While it relied on the measurement of carbon dioxide respiration to determine microbial activity, another study concluded that freezing had no significant effect on the microbial activity in surrounding burial soil (Stokes et al., 2009).

Nonetheless, it is essential to also examine any freezing effects immediately after freeze-thaw process and prior to decomposition. This is especially important after the publication of a case study that showed a shift in post-mortem microbial community abundance and structure during the thawing of two pediatric cases (Pechal et al., 2017). While the sample size for the study was limited, their observation called for an in-depth look at the effects of freezing donors and what it might mean for microbiome research in forensic taphonomy.

The purpose of this study therefore was to determine if the storage of donors below freezing at human taphonomy facilities may have any effect on their microbiome. Seven donated bodies at the Forensic Anthropology Center at Texas State (FACTS) were swabbed before they were frozen and after the complete thawing of the bodies. It was also assessed if the freezing of the bodies may have varying effects at different anatomical locations (n = 5), and how the microbial composition at these locations may differ from one another post-mortem.

Materials and methods

Sampling

This research was organized and carried out in collaboration with FACTS as a part of the “Mass Grave Project,” an interdisciplinary taphonomic research project aimed at improving the detection, excavation and documentation of clandestine mass and single graves (Mickleburgh et al., 2022). Whole body donations studied at FACTS are acquired for scientific research purposes, through the expressed and documented willing of the donors and/or their legal next of kin. All human decedents involved in this study were collected through a body donation program which complies with all legal and ethical standards associated with the use of human remains for scientific research in the USA (Gocha et al., 2021). The sampling of tissues from the body donors in the Mass Grave Project was conducted in accordance with the Texas Uniform Anatomical Gift Act.¹ Ethical approval for analysis of the samples at Northumbria University was provided by the Northumbria ethics committee (ref: 21514). Nine body donations were received for the Mass Grave Project, however, microbiome analysis for the present study was conducted on only seven of these donors for which both pre- and post-freezing samples were available.

Initial samples were collected upon arrival of the donors at the research facility, either immediately or after a period during which bodies were kept refrigerated (4°C) due to technical reasons (Table 1). Swab samples, collected in triplicates, were taken from five anatomical locations of each corpse. Areas sampled included hand, foot, neck, oral cavity, and rectum.

TABLE 1

Table 1. Duration of refrigeration (+4°C) between arrival to the facility and swabbing, between swabbing and freezing of the donors, and duration of freezing (−20°C) of the donors.

Hand and foot samples were taken from the dominant limb as identified by the donor information provided. Swabs were collected using sterile swabs by rubbing the swab at the sampling location (i.e., hand, foot, neck, oral cavity, or rectum) for 15 or 30 s, depending on the location. Hand or foot swabs sampled the palm and fingers or sole and toes respectively for 15 s. Neck samples were taken at the back, in the area between the base of the head and the top of the shoulder, by swabbing back-and-forth for 15 s.

For the oral cavity, samples were collected from the left cheek, right cheek, and oral vestibule, including any teeth if available, by swabbing the aforementioned areas using the same swab for 15 s. Finally, rectal samples were obtained by inserting the swab into the anus to about 5 – 10 cm into the rectum, and then rotating for 30 s.

After initial sampling as described, swab samples were stored immediately at −20°C. Bodies were frozen either immediately post-swabbing, or after a short period of time during which they were kept refrigerated (Table 1) due to technical reasons. Due to the differing arrival times of the donors to the facility, the total time spent frozen in storage varied among them. However, all bodies remained in storage at −20°C for a minimum of period of 11 days and a maximum of 40 days (Table 1). For ease of reference, all donors were assigned a reference label – D12, D13, D14, D20, D21, D22, and D23.

After the freezing period, donors were removed from storage and thawed. Donor bodies were frozen and defrosted within a closed and controlled indoor laboratory (21°C) to prevent insect access. Thawing began at about 7:00 pm on the 10th of May 2021, and post-freezing sampling commenced on the 12th of May at about 8:30 am, after complete thawing of the bodies. Swab samples were taken again from each body, and location, as previously described. All swabs were immediately stored frozen at −20°C, along with samples taken prior to freezing. All samples were then shipped to Northumbria University for processing.

DNA extraction

Microbial DNA was isolated from the swab samples using the DNeasy 96 PowerSoil Pro Kit from Qiagen (Qiagen, Hilden, Germany) with a modified protocol. 800 uL of Solution CD4 was added to Eppendorf tubes containing swab samples, vortexed for 30 s and centrifuged at 5000 rpm for 10 min. The supernatant was then transferred to the PowerBead Pro Plate, after which the extraction was performed according to the manufacturer’s protocol.

In addition to swab samples, two extraction negatives – one with a sterile swab, and the other using just kit reagents – were included to identify any potential contamination. A positive swab of a bacterial culture – E. coli Top10 plo3 D55N Hox A – was also used to check the quality of microbial DNA extraction.

Quality check analysis was further analyzed qualitatively using PCR products of extracted microbial DNA via gel electrophoresis. This included the extraction controls (negatives – sterile swab, “N” and only kit reagents, “N1” – and positive, “P”) and a few randomly selected samples from the experiment (Supplementary Figure 1). PCR reaction mixtures for the quality check were set up following Procopio et al. (2021) using 2.0 μL template DNA in a final reaction volume of 25 μL. 1.5% (w/v) agarose gel made with 1× TAE buffer was used to check quality of PCR products obtained prior to library preparation and sequencing.

Sequencing and bioinformatic analysis

Library preparation and sequencing of the 16S rRNA gene (V4 region) was analyzed by NU-OMICS (Northumbria University, Newcastle, UK) based on the Schloss wet-lab MiSeq SOP (Kozich et al., 2013). Briefly, PCR was carried out using 1× Accuprime Pfx Supermix, 0.5 μM each primer and 1 μL of template DNA under the following conditions 95°C 2 min, 30 cycles 95°C 20 s, 55°C 15 s, 72°C 5 min with a final extension 72°C 10 min. One positive and one negative control sample were included in each 96 well plate and carried through to sequencing.

PCR products were normalized using SequalPrep™ Normalization kit (Invitrogen) as described in the manufacturer’s instructions and combined into four pools. Each pool was quantified using fragment size determined by BioAnalyzer (Agilent Technologies) and concentration by Qubit (Invitrogen). Pools were combined in equimolar amounts to create a single library then denatured using 0.2N NaOH for 5 min and diluted to a final concentration of 5 pM, supplemented with 15% PhiX and loaded onto a MiSeq V2 500 cycle cartridge (Illumina Inc., San Diego, CA, USA). Targeting and sequencing of the V4 region of the 16S rRNA for bacterial identification were done following the gold standards suggested by the Human Microbiome Project.

Fastq raw sequencing files generated were imported into QIIME2 software ver. 2021.11 for analysis and quality filtered using DADA2 and trimmed at 240 bp forward and 222 bp reverse. Taxonomy was assigned using the naive Bayes classifier, which was trained on silva-138-99 operational taxonomic units (OTUs). Qiime2 output artifacts were analyzed in R (ver. 4.2.2) using the phyloseq package. Abundance and diversity metrics calculated level were assessed by the aggregation of taxa to phylum or genus level. At genus level, unassigned taxa labeled “NA,” “unassigned,” “metagenome,” “uncultured” were excluded to prevent problems in agglomeration due to multiple taxa being assigned the same name.

Results

A total of 35 pairs of swab samples, taken from seven donors at five anatomical locations – hand, neck, foot, oral and rectum – were analyzed in this study. Each pair was composed by a swab sample taken before freezing (“fresh”), and another taken after freezing the body at −20°C (“thawed”). Qualitative analysis to assess successful extraction and amplification of the 16S rRNA gene using gel electrophoresis (Supplementary Figure 1) showed no amplification signals in both negative controls, a strong band for the positive control, and positive amplification for all samples, although one displayed a weak signal.

Following extraction, amplification and sequencing, the resulting data were standardized to the median sampling depth to account for unequal sequencing depth. This allowed inclusion of all samples to prevent waste of data. The Shannon index – accounting for microbial richness and evenness – was used to determine if there were any differences in the alpha diversity of the bacterial communities within the fresh and thawed samples. No significant differences were observed overall between all fresh and thawed samples (Wilcoxon; p = 0.48). Furthermore, analysis of the alpha diversity, also using the Shannon index, between fresh and thawed samples at each sample source (hand, neck, foot, oral and rectum), showed no significant differences (Figure 1).

FIGURE 1

Figure 1. Alpha diversity plot (Shannon diversity) of fresh and thawed samples per anatomical location at ASV level.

The abundance distribution of the bacterial communities in fresh and thawed samples at phylum level was evaluated to detect any microbial compositional changes resulting from the freezing of the donors. Overall, statistical analysis showed that phylum abundances between the two groups were not significantly different. However, Bacteroidota and Firmicutes abundance seemed to decrease in thawed samples, while Proteobacteria appeared to increase after freezing (Figure 2A). At genus level, five taxa Christensenella, Anaerotruncus, Carnobacterium (phylum – Proteobacteria), Acinetobacter, Psychrobacter (phylum – Firmicutes) had significantly different abundances across the two sample groups (Figure 2B). Finally, two phyla, Deinococcota and Myxococcota, were only found in thawed samples, albeit at low abundance levels.

FIGURE 2

Figure 2. Abundance plot of fresh and thawed samples at phylum level (A) and taxa at genus level (B) with abundance significantly different across fresh and thawed samples.

Shannon diversity of the samples by anatomical location at ASV level, disregarding when they were collected (i.e., before or after freezing), showed that rectum samples were significantly different to other samples taken from any other location (Figure 3). Additionally, we explored sample distribution with Principal Coordinates Analysis (PCoA), using Bray–Curtis distance at ASV level to investigate any inter-individual microbial variation in relation to the sample source or anatomical location and when the samples were collected (Figures 4A, B). There were no observable clusters of fresh or thawed samples, or samples taken from hand, neck, foot, or oral locations. An ADONIS test (with vegan ver. 2.6-2) using Bray–Curtis distance measurement also showed no significant difference at genus level between fresh and thawed samples (p = 0.7439; permutation = 10000; seed = 22).

FIGURE 3

Figure 3. Violin plot showing Shannon diversity at each anatomical location at ASV level. * less than or equal to 0.01; ** less than or equal to 0.001; *** less than or equal to 0.0001.

FIGURE 4

Figure 4. Bray–Curtis distance plot of samples at each anatomical location (A) and plots showing shift in donor microbial community after freezing at each anatomical location (B).

However, there was a clear separation of samples taken from the rectum, reflecting the significant difference in alpha diversity observed in Figure 3. Interestingly, all samples, including rectum samples, collected from one donated donor, D22, were also well separated from all the remaining donors.

Discussion and conclusion

The practice of freezing human bodies as a means of preserving the composition and abundance of associated microbial communities as seen ante-mortem or prior to experimental research is not unusual, especially at human taphonomy facilities (Hyde et al., 2013, 2015). It has been shown that microbial signatures can be obtained from human DNA extracts up to 16 years of being frozen in storage (Sguazzi et al., 2022). However, reports on how freezing human-related samples affects the microbiome has been varied. More specifically, how freezing the entire body affects their microbiome is unknown. Our analysis of seven donors at FACTS facility showed that overall, storing bodies frozen preserves the composition and abundance of their microbiome. Our results also demonstrate that bacterial community composition in the rectum post-mortem is significantly different to other anatomical areas, regardless of freezing.

Observing similar microbiome signatures in the donors before and after freezing is unsurprising as it is generally accepted that for a variety of human-related samples – including fecal matter (Cardona et al., 2012; Pérez-Burillo et al., 2021), skin and saliva (Marotz et al., 2021; Furuhashi et al., 2022), meconium (Xin et al., 2021), and milk (Lyons et al., 2021) – freezing is sufficient to preserve microbial composition. This is supported by the universal knowledge that low temperatures can slow or stop the proliferation of bacteria already present within samples and hinder the establishment of potential contaminants. Hence, the initial baseline composition and abundance of microbial communities in the sample is preserved.

However, although classified as statistically non-significant, our results still show that there were changes to the microbiome after freezing. Pérez-Burillo et al. (2021) have speculated that certain taxa may become impaired due to freezing, allowing competing taxa to thrive in their absence after thawingClick or tap here to enter text.. This change in ecological relations may explain the presence or absence of taxa, such as Deinococcota and Myxococcota phyla, despite the overall stability of the microbiome after freezing. Such observations are important to note, particularly for the interpretation of microbiome research in forensics, as oftentimes “rare” taxa can be presented as trace evidence (Young and Linacre, 2021) or potential biomarkers in human profiling (Hampton-Marcell et al., 2020; Neckovic et al., 2020).

The outcome of this research also provided insights into the post-mortem microbial community at various locations of the human body. Generally, the rectal, oral, and skin microbiomes in life have been reported to differ significantly in their community composition (Costello et al., 2009; Huttenhower et al., 2012). Our results, however, show that apart from the rectum, microbial communities in these habitats are hardly distinguishable after death. While microbial community homogenization occurs after death (Burcham et al., 2019; Liu et al., 2021), particularly with post-mortem transmigration (Burcham et al., 2016), this is unlikely in this case as the donors were stored frozen and we have shown that this storage method does not significantly change the microbiome composition. One thing to note is that all sites sampled – excluding rectum and oral – were all skin sites and this might contribute to the similarity observed. We also propose that this similarity may be caused by the fact that the microbial community differences between oral and skin habitats are not large enough to be detected with our limited sample size. This is supported by another study reporting that communities in various human body habitats are distinct in the post-mortem human microbiome (Pechal et al., 2018). Additionally, the rectal microbiome recorded the smallest shift after freezing and thawing (Figure 4B). This may indicate that the rectal microbiome was the least affected by freezing and is therefore more stable. The sample size included in this study is however not enough to conclusively determine this point.

Furthermore, PCoA grouping of samples (Figure 4A) revealed a deviation in the behavior of rectal samples from one donor, D22, compared to other rectal samples. Closer examination of the plot showed that all samples collected from this donor clustered closely together, regardless of the anatomical origin of the sample. It is unclear why this occurred, although, medical records for this donor were extensive. One thing of note in their record was their gastrostomy status which showed that this donor had a feeding tube antemortem. Studies have shown that enteral feeding tubes can cause alterations in the human microbiome, particularly oral (Takeshita et al., 2011) and gut (Whelan et al., 2009; Fan and Lee, 2021) microbiota. More importantly however, despite the reasons for this alteration, our study shows that complex or deviant microbiomes of donors – even those with complicated medical histories antemortem – can be preserved by freezing, as the microbial shift in D22’s samples prior to and after freezing was not significant.

It is important to note however that this study did not assess the viability of these bacteria as only the genetic material – the microbiome – was evaluated. Research assessing the survival of microorganisms in frozen foods (Speck and Ray, 1977) however, has shown that these microbes can continue to persist, and function. Moreso, another study indicated that lesions in microbial DNA can be repaired at −15°C (Dieser et al., 2013). Therefore, we conclude that freezing human donors at −20°C does not introduce a substantial alteration in their associated microbiome signatures. While we observed small changes to their microbiomes, these changes were not statistically significant to warrant an overhaul in the storage method of donors, particularly at forensic taphonomy facilities when performing thanatomicrobiome research.

Data availability statement

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found below: https://www.ncbi.nlm.nih.gov/, PRJNA925138.

Ethics statement

This study did not involve living human subjects but included only human tissues from deceased individuals from which informed consent was obtained prior to death or from legal next of kin. The study is not subject to institutional review. The ethics code of FACTS was followed and approval for the study was obtained from the FACTS Ethics Review Board as well as from the Ethics Committee at Northumbria University Ref. 21514.

Author contributions

NP, SG, and HM contributed to the conception and design of the study. NP, HM, and TG performed the samplings. NO and AN conducted the laboratory research. NO performed the statistical analysis reviewed by AN. NO wrote the first draft of the manuscript. NP, DW, and HM administered the study. All authors contributed to the manuscript revision, read, and approved the submitted version.

Funding

The mass grave project was supported by the VINNOVA – funded research project “THANATOS” (HM) under grant 2019–05211 and by the UKRI Future Leaders Fellowship (NP) under grant MR/S032878/1.

Acknowledgments

We thank Prof. Darren Smith and Stefano Ghignone for their insightful comments and suggestions regarding statistical analysis.

Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Publisher’s note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fevo.2023.1151001/full#supplementary-material

Supplementary Figure 1 | Gel electrophoresis of randomly selected samples after DNA extraction and PCR amplification with positive (P) and negative (N and N1) extraction controls. Sample names are listed by donor ID (Dx), anatomical location (H – hand, N – neck, F – foot, O – oral, and R – rectum) and freezing status (F – fresh and T – thawed).

Footnotes

1. ^ https://statutes.capitol.texas.gov/Docs/HS/htm/HS.692A.htm

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