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 .