Abstract
Encapsulated systems have been widely used in environmental applications to selectively retain and protect microorganisms. The permeable matrix used for encapsulation, however, limits the accessibility of existing analytical methods to study the behavior of the encapsulated microorganisms. Here, we present a novel method that overcomes these limitations and enables direct observation and enumeration of encapsulated microbial colonies over a range of spatial and temporal scales. The method involves embedding, cross-sectioning, and analyzing the system via fluorescence in situ hybridization, while retaining the structure of encapsulants and the morphology of encapsulated colonies. The major novelty of this method lies in its ability to distinguish between and subsequently analyze multiple microorganisms within a single encapsulation matrix across depth. Our results demonstrated the applicability and repeatability of this method with alginate-encapsulated pure (Nitrosomonas europaea ) and enrichment cultures (anammox enrichment). The use of this method can potentially reveal interactions between encapsulated microorganisms and their surrounding matrix, as well as quantitatively validate predictions from mathematical models, thereby advancing our understanding of microbial ecology in encapsulated, or even biofilm systems, and facilitating the optimization of these systems.
Introduction
Microbial encapsulation is a versatile environmental biotechnology with applications ranging from nutrient removal and resource recovery from wastewater to bioremediation of contaminants . The permeable matrices used as encapsulants not only selectively retain microorganisms, but also protect them against potential inhibitors in the external environment . Nevertheless, encapsulants create physical barriers to directly access microbes for analysis. Existing tools have diverse capabilities in microbial and morphological analysis but are limited when applied to elucidate the growth of multiple encapsulated microorganisms across a temporal and spatial span.
Due to the similarity between biofilm and encapsulated systems, microscopic methods are widely used in both systems to reveal the morphology and assembly of microbes. For instance, optical microscopy, optical coherence tomography, and electron microscopy have been used to reveal structures in biofilm systems , which are applicable to encapsulated systems as well. Optical and fluorescent dyes can be used to enhance the contrast between the encapsulant and encapsulated microorganisms . Live/dead staining is also available to identify living encapsulated cells . Meanwhile, encapsulants are inherently different from extracellular polymeric substances in biofilms. It is challenging to preserve their structure during the pretreatment of samples. To prepare samples for microscopy, fixation and dehydration procedures can cause shrinkage and even destruction of hydrogel-derived encapsulants. . Electron microscopy can better preserve encapsulant structures and improve the resolution of submicron morphological details; it cannot, however, differentiate between microbial species.
In addition to microscopic methods, molecular and physio-chemical techniques are used to quantify the growth of encapsulated microorganisms. Measurable parameters include culturable colony forming units (CFU), DNA copies, cell protein content, and dry cell weight . Sequencing and quantitative amplification (e.g., quantitative polymerase chain reaction (qPCR)) of genetic material can further identify and quantify the targeted components of an encapsulated microbial community . Nevertheless, extraction of cell contents or DNA from encapsulated microorganisms often requires homogenization and removal of the encapsulant. As a result, the spatial information of cell distribution within the encapsulant is lost during such a process. Modifications of the procedure with selective shearing of surface layers and sorting of size fractions have been demonstrated with granular activated sludge; however, these modifications are difficult to apply on encapsulated systems, as encapsulants are much harder to selectively disassemble.
The main objective of this study was to develop a novel method for measuring spatial and temporal changes in the encapsulated growth of various microorganisms. Such a method can potentially provide information to validate predictions from mathematical models, allowing more predictable applications of microbial encapsulation technologies in environmental engineering .
Methods and Materials
2.1 Microbial encapsulation and batch test setup
A pure culture of Nitrosomonas europaea ATCC 19718T and an enrichment culture of anammox bacteria were used as inocula for microbial encapsulation. The enrichment culture was granular in nature and collected from a bench-scale sequencing batch reactor under steady-state operation . An equal volume of 4% sodium alginate solution and the inoculum culture was completely mixed before crosslinking in a 4% calcium chloride solution to form spherical beads as described previously (Wang et al., 2022). The inocula were about 20 and 500 mgTSS/L for the Nitrosomonas culture and anammox enrichment respectively. Crosslinked alginate beads had diameters of 2.4 – 2.8 mm.
Batch bioreactors with encapsulated biomass were established as previously reported . For tests with encapsulated Nitrosomonas , ammonia (50 – 150 mgN/L) was added daily to the synthetic culture medium as NH4Cl, whereas an equal amount of ammonia (as NH4Cl) and nitrite as NaNO2 (50 – 100 mgN/L each) was added to the medium for tests with the anammox enrichment culture. Triplicate batch reactors (200 mL) were placed on an orbital shaker at 100 rpm and operated for a week. Sponge caps were used to allow diffusion of air into the reactors, and dissolved oxygen concentrations were kept between 0.5 to 4.0 mg/L. Water samples (1.5 mL) were collected daily, filtered, and analyzed with spectrophotometric methods for ammonia, nitrite, and nitrate . Six beads were sampled (3 for imaging and 3 for DNA extraction) and processed immediately before and after each batch incubation.
2.2 Sample preparation, cross-sectioning, and FISH procedure
Alginate bead samples were fixed immediately after collection with 3% (w/v) paraformaldehyde. Each alginate bead was placed in the conical bottom of a 1.5-mL microcentrifuge tube and infiltrated stepwise with 25, 50, 75, and 100% glycolmethacrylate (GMA) as previously described . GMA-infused samples were thus molded into the tip of the microcentrifuge tube. Following the solidification of GMA, the bottom of the microcentrifuge tubes was cut open and placed on a microtome (Microm HM505E). Cross-section cutting (10-µm thickness) was conducted starting from the surface of each bead. Three consecutive sections were collected upon every 100-µm advancement until the entire bead was completely cross-sectioned (Figure 1 ). The collected sections were placed on multi-well slides pre-coated with a 0.1% poly-L-lysine solution. Fluorescence in situ hybridization (FISH) was conducted with oligonucleotide probes targeting ammonia oxidizing bacteria (AOB), nitrite oxidizing bacteria (NOB), and anammox bacteria (Table 1 ). The hybridization was conducted with 35% formamide at 46 °C for 2 hours. An antifade mountant with DAPI (ProLong™ Gold, Invitrogen) was used as a DNA counter stain. Alginate beads without microbial inoculum were analyzed by the cross-section FISH method as negative controls using each fluorescence probe to check for autofluorescence. The anammox enrichment culture was used as a positive control for each fluorescence probe, as it was assumed to contain AOB, NOB, and anammox bacteria. Prior to fixation, the granular anammox culture was homogenized for one minute with a handheld homogenizer to break apart large granules. The anammox biomass was then fixed and used for FISH as described above, without the embedding material present. GMA has been previously shown to not interfere with, but rather, to improve the sensitivity of FISH analysis .