김수빈
(Subin Kim)
aiD
김상균
(Sangkyun Kim)
biD
박면호
(Myeonho Park)
ciD
김성아
(Seunga Kim)
diD
이민주
(Minjoo Lee)
eiD
박준홍
(Joonhong Park)
f†iD
-
연세대학교 공과대학 건설환경공학과
(Department of Civil and Environmental Engineering, Yonsei University)
Copyright © KOREAN SOCIETY ON WATER ENVIRONMENT
Key words
Anammox bacteria, Freeze-thaw cycle, Low-temperature acclimation, Specific anammox activity, Nitrogen removal, Water quality
1. Introduction
Anaerobic Ammonium Oxidation (Anammox) bacteria play a critical role in the nitrogen
cycle by converting fixed nitrogen species, such as ammonia (NH₄⁺) and nitrite (NO₂⁻),
into nitrogen gas (N₂) under anaerobic conditions(Mulder et al., 1995; Strous, Fuerst et al., 1999; Van de Graaf et al., 1995). Wastewater treatment using Anammox is highly energy-efficient compared to conventional
nitrogen removal methods and does not require additional organic carbon sources, making
it particularly advantageous in terms of environmental sustainability and cost-effectiveness(Abma et al., 2010; Kartal et al., 2010). However, the application of Anammox technology remains largely limited to sidestream
wastewater treatment with high nitrogen concentrations, and its implementation in
mainstream wastewater treatment is still under investigation(van der Star et al., 2007). One of the primary challenges for mainstream applications is the temperature sensitivity
of Anammox bacteria. Their optimal activity range lies between 30°C and 40°C, and
metabolic activity declines sharply at lower temperature (8-15°C), significantly reducing
specific Anammox activity (SAA)(Lotti et al., 2015). This temperature dependency poses a major constraint for applying Anammox processes
in regions with seasonal temperature variations.
Despite these limitations, the potential for mainstream application remains promising
because of the ecological significance of Anammox bacteria. These microorganisms are
found in diverse environments, such as freshwater, marine ecosystems, soil, and rice
paddies, and can survive across a broad temperature range of -30°C to 80°C, contributing
substantially to global nitrogen(Oshiki et al., 2016; Wang et al., 2019). Anammox bacteria have been shown to actively contribute to nitrogen removal in extreme
environments, such as the Arctic Ocean, where they maintain high activity even at
-1.3°C(Hu et al., 2013; Oshiki et al., 2016; Rysgaard and Glud, 2004; Rysgaard et al., 2004). Furthermore, species such as Candidatus Brocadia and Candidatus Jettenia have been
detected in extreme freshwater environments below -30°C, demonstrating their adaptability(Zhu et al., 2015). Notably, Candidatus Scalindua is known for its activity in cold marine environments,
where it plays a critical role in global nitrogen cycling(Dalsgaard and Thamdrup, 2002). The presence of such species suggests that Anammox bacteria have evolved through
selective growth, adaptation, and mutation, enabling their survival across diverse
and extreme environments.
Studies on evolutionary adaptation have indicated that bacteria exposed to environmental
stress undergo selective growth and metabolic changes to enhance their survival (Berry and Foegeding, 1997; Fuerst and Sagulenko, 2011; Sharma et al., 2006). The first stage of this process involves the selective enrichment of specific populations,
which can lead to acclimatization and genetic mutations. Bennett and Lenski’s experiments
with E. coli under varying thermal conditions provide evidence of rapid evolutionary
adaptation in stressful environments, highlighting trade-offs in temperature-dependent
fitness(Bennett and Lenski, 1993, 1997, 1999; Lenski, 2017; Parsons, 1987). These findings suggest that specific populations of microorganisms can undergo selective
enrichment under environmental stress, leading to adaptation and evolutionary shifts
in the community structure. The observed adaptability of Anammox bacteria in diverse
ecosystems supports this hypothesis, indicating the need to evaluate their potential
for selective enrichment and survival under extreme conditions in engineered environments.
Research on the low-temperature adaptation of Anammox bacteria has focused on two
primary areas. The first explores strategies for adaptation to low temperatures (<20°C)(De Cocker et al., 2018; Fukunaga et al., 1999; Hendrickx et al., 2014; Kouba et al., 2022; Wu et al., 2016). The second study investigated preservation strategies to maintain bacterial viability
during Freeze-Thaw Cycles (FTC) and assessed structural changes to identify optimal
preservation conditions. Previous studies have demonstrated high reactivation rates
under cryogenic conditions at -80 and -200°C(Ali et al., 2014; Heylen et al., 2012; Huang et al., 2022; Ji and Jin, 2014; Rothrock et al., 2011). Despite evidence of low-temperature adaptation, studies on selective enrichment
under stressful conditions are limited. By applying the principles of evolutionary
adaptation, the selective enrichment of specific populations can be promoted, thereby
increasing the likelihood of survival in cold environments. FTC are critical environmental
stressors that promote the selective enrichment of resilient Anammox bacteria. Prior
studies on microorganisms, such as E. coli, have shown enhanced survival and activity
under repeated FTC exposure(Sawicka et al., 2010; Sleight and Lenski, 2007). These findings highlight the potential of stress-induced adaptation to enhance initial
survival and efficiency in extreme environments, ultimately supporting nitrogen removal
at low temperatures.
This study aimed to investigate the effect of repetitive freeze-thaw stress pre-treatment
on the adaptation of Anammox bacteria during low-temperature reactivation by evaluating
their survival and functional recovery with a focus on nitrogen removal efficiency
and metabolic activity. Specifically, this study assessed whether Anammox bacteria
can undergo selective enrichment and adapt to low-temperature environments within
wastewater treatment systems, similar to their behavior in natural ecosystems, in
order to elucidate their biological adaptation mechanisms. To this end, repeated freeze-thaw
cycles were applied under cryogenic conditions (-80°C and -200°C), which have previously
been shown to achieve high reactivation rates(Ali et al., 2014; Chen and Jin, 2017; Heylen et al., 2012; Rothrock et al., 2011). After inducing selective enrichment and initial adaptation through the FTC, the
reactivation performance under low-temperature conditions was evaluated. This study
provides fundamental insights to enhance the applicability of Anammox bacteria in
mainstream wastewater treatment processes during winter conditions.
2. Materials and Methods
2.1 Inoculum source
The Anammox inoculum used in this study was sourced from a continuously stirred tank
reactor (CSTR) that had been operated for one year. The initial biomass concentration
was 14,000 mg-MLSS/L. To prevent oxygen from inhibiting Anammox activity, the reactor
was maintained under anaerobic conditions, ensuring that the dissolved oxygen (DO)
level remained below 0.2 mg/L. The operational temperature was maintained at 30 ±
2°C. Ammonium chloride (NH₄Cl) and sodium nitrite (NaNO₂) were supplied as nitrogen
sources, resulting in a total nitrogen (TN) concentration of 264 mg-N/L in the feed
solution. The reactor consistently achieved a nitrogen removal efficiency (NRE) of
over 95% with pH controlled at 7.5 ± 0.3. Microbial community analysis revealed that
Candidatus_Kuenenia, Candidatus_Brocadia, and Candidatus_Brocadiaceae_unclassified
were the dominant genera, collectively accounting for > 23% of the total microbial
population.
2.2 Reactivation strategy
To evaluate the impact of FTC on the recovery of Anammox bacteria, three cycles were
conducted, each followed by a reactivation phase(Fig. 1). A control group, referred to as No Freeze-Thaw (NFT), was also included. This group
was not subjected to any freeze-thaw treatment but was incubated under the same reactivation
conditions (15°C), allowing for direct comparison with FTC-treated groups. The samples
were frozen at -80°C and -200°C and subsequently reactivated at 15°C for five days.
Biomass was obtained from the parent culture and standardized to 5,000 mg-MLSS/L to
ensure uniformity across all test groups. All experimental steps were performed in
an anaerobic chamber to limit exposure to dissolved oxygen level. The FTC process
spanned nine days, with each cycle lasting for three days. Prior to freezing, residual
substrates were removed by rinsing the Anammox biomass three times with 0.1 M phosphate
buffer (pH 7.2). The prepared samples were maintained at -80°C in deep freezers and
at -200°C in liquid nitrogen. After completing one, two, or three cycles, the biomass
was transferred to 160 mL serum bottles with an operational volume of 100 mL and incubated
at 15°C for reactivation. The reactivation phase was conducted using a Sequencing
Batch Reactor (SBR) system. The SBR operation cycle included a 30-minute filling stage,
a 22-hour reaction stage, a one-hour settling stage, and a 30-minute drawing stage,
yielding a Hydraulic Retention Time (HRT) of 24 hours. On the fifth day of reactivation,
both biological and chemical analyses were conducted to assess the adaptation and
recovery of the Anammox bacteria.
Fig. 1. Reactivation protocol.
2.3 Synthetic wastewater
The synthetic wastewater used in this study was prepared based on the mineral medium
composition listed in Table 1. Ammonium chloride (NH₄Cl) and sodium nitrite (NaNO₂) were used as nitrogen sources,
with final concentrations of 60 mg-N/L and 72 mg-N/L, respectively. To stabilize the
alkalinity, 5 mM sodium bicarbonate (NaHCO₃) was added as a buffering agent. Essential
nutrients, including KH₂PO₄ (27 mg/L), MgSO₄⋅7H₂O (300 mg/L), and CaCl₂⋅2H₂O (180
mg/L), were incorporated to support bacterial growth. The medium was supplemented
with two distinct trace-element solutions. Trace element solution I provided essential
iron through 5000 mg/L of ethylenediaminetetraacetic acid (EDTA) and 5000 mg/L of
FeSO₄⋅7H₂O. Trace element solution II contained EDTA (15,000 mg/L) and other trace
elements critical for metabolic activity and bacterial proliferation: ZnSO₄⋅7H₂O (430
mg/L), CoCl₂⋅6H₂O (240 mg/L), MnCl₂⋅4H₂O (990 mg/L), CuSO₄⋅5H₂O (250 mg/L), Na₂MoO₄⋅2H₂O
(220 mg/L), NiCl₂⋅6H₂O (190 mg/L), Na₂SeO₄⋅10H₂O (210 mg/L), and H₃BO₃ (14 mg/L).
This medium composition provided all essential nutrients and trace elements required
to support bacterial activity and growth.
Table 1 Wastewater quality of mineral medium
|
Medium
|
1 L
|
|
NH4Cl (60 mgN/L, 99%)
NaNO2 (72 mgN/L, 98%)
NaHCO3 (5 mM)
KH2PO4
MgSO4⋅7H2O
CaCl2⋅2H2O
|
0.232 g
0.362 g
0.42 g
0.027 g
0.3 g
0.18 g
|
|
Trace
elemental
solution Ⅰ
|
DI
EDTA
FeSO4
|
1 L
5 g
5 g
|
|
Trace
elemental
solution Ⅱ
|
DI
EDTA
ZnSO4⋅7H2O
CoCl2⋅6H2O
MnCl2⋅4H2O
CuSO4⋅5H2O
NaMoO4⋅2H2O
NiCl2⋅6H2O
NaSeO4⋅10H2O
H3BO4
|
1 L
15 g
0.43 g
0.24 g
0.99 g
0.25 g
0.22 g
0.19 g
0.21 g
0.014 g
|
2.4 Chemical and physical analysis
Samples were collected daily during the 5-day reactivation period using a fill-and-draw
process, in which the medium was replaced with fresh substrate solution each day to
maintain consistent anaerobic conditions and nutrient supply. The collected samples
were then filtered through a 0.45-μm syringe filter, and promptly analyzed to determine
the concentrations of ammonium nitrogen (NH₄⁺-N), nitrite nitrogen (NO₂⁻-N), and nitrate
nitrogen (NO₃⁻-N). The concentration of ammonium nitrogen (NH₄⁺-N) was quantified
using the Salicylate Method(Hach Method 10023), with a detection range of 0.02–2.5
mg/L at a wavelength of 342 nm. nitrite nitrogen (NO₂⁻-N) was measured following the
Standard Method 4500-NO₂-B(Colorimetric Method), offering a detection limit of 0.4
mg/L at 543 nm. Nitrate nitrogen (NO₃⁻-N) was analyzed using the Chromotropic Acid
Method(Hach Method 10020), which has a detection range of 0.2–30 mg/L at 344 nm. The
pH levels and dissolved oxygen (DO) concentrations were monitored using a Thermo Orion
Star A3295 pH/DO meter. All measurements were conducted in duplicate to ensure accuracy,
and mean values were reported.
2.4.1 Specific Anammox activity
Specific Anammox activity (SAA) was evaluated through experiments conducted using
a sequencing batch reactor (SBR) during a short-term study. Biomass samples of Anammox
sludge were collected following each freeze-thaw cycle (1FTC, 2FTC, and 3FTC). The
harvested biomass was rinsed three times with 0.1 M phosphate buffer at pH 7.2 to
remove residual substrates. The SBR system utilized in the experiment had an operational
capacity of 100 mL, and the biomass concentration was standardized to 4.014 kg-VSS/m³.
The initial concentrations were set to 60 mg-N/L for ammonium nitrogen (NH₄⁺-N) and
72 mg-N/L for nitrite nitrogen (NO₂⁻-N). To create anaerobic conditions, the reactor
was purged with 99.99% pure nitrogen gas for 10 min and was securely sealed. The SBR
process operated in distinct cycles, including filling, reaction, settling, and decanting
phases, with all steps performed under carefully controlled conditions. The reaction
phase was conducted in a thermostatic shaker maintained at 15 ± 1°C and set to a stirring
speed of 200 rpm. During this phase, hourly sampling was performed to monitor the
reduction in ammonium nitrogen (NH₄⁻-N) and nitrite nitrogen (NO₂⁻-N) concentrations.
The SAA was calculated by determining the nitrogen removal rate (NRR) and normalizing
it against the volatile suspended solid (VSS) concentration. The results were expressed
as kg-N/kg-VSS per day (d). All experiments were conducted in duplicate to ensure
accuracy, and average values were used for analysis. At the end of each cycle, sludge
samples were collected for further investigation of the microbial community structure
and molecular properties.
2.5 DNA/RNA extraction
Genomic DNA was isolated from the samples using the DNeasy PowerSoil Kit(Qiagen, Carlsbad,
CA, USA) following the manufacturer’s protocol. A maximum of 0.25 mg of the sample
was added to a PowerBead Pro Tube containing 800 µL of Solution CD1 for cell lysis.
The mixture was vortexed for 10 min to ensure complete disruption of the cells. Following
vortexing, the mixture was centrifuged at 15,000 × g for 1 min and the supernatant
was carefully transferred to a new tube. Subsequently, 200 µL of Solution CD2 was
added, followed by vortexing and centrifugation to remove the inhibitors from the
supernatant and discard the supernatant. The supernatant was collected again and mixed
with 600 µL of Solution CD3 to bind the nucleic acids. The resulting lysate was applied
to an MB Spin Column, which was centrifuged to isolate the DNA. The column was sequentially
washed with Solutions EA and C5 to remove impurities such as proteins and other contaminants.
Finally, the DNA was eluted with 60 µL of Solution C6 (nuclease-free water). DNA concentration
was measured using a NanoDrop Spectrophotometer(Thermo Scientific, Waltham, MA, USA),
and the concentration was adjusted to approximately 2 ng/µL. All DNA samples were
stored at -20°C until further analysis.
To prevent RNase contamination, RNA extraction was performed under strict RNase-free
conditions. The SPINeasy RNA Kit(MP Biomedicals, Irvine, CA, USA) was used according
to the manufacturer’s protocol. To collect the cell pellet, 0.25 mL of bacterial culture
was centrifuged at 10,000 × g for 3 min. The resulting pellet was resuspended in 250
µL RNASS, transferred into a Lysing Matrix B tube, and mixed with 750 µL Lysis Buffer
R. Cell lysis and homogenization were achieved using a FastPrep instrument at a speed
of 6.0 m/s for 40 s. Following centrifugation of the lysate at 14,000 × g for 10 min,
the supernatant was combined with an equal volume of ethanol and applied onto a spin
column. To eliminate any residual DNA, DNase I treatment was applied directly to the
column membrane for 15 min at RT. The column was washed twice with Wash Buffer R to
remove impurities, and the RNA was eluted in 60 µL of nuclease-free water. RNA concentration
was measured using a NanoDrop Spectrophotometer(Thermo Scientific, Waltham, MA, USA)
and calibrated to approximately 10 ng/µL. The extracted RNA was stored at -80°C for
future analyses.
2.6 Quantitative real-time PCR (qPCR)
Bacterial biomass was quantified using quantitative real-time PCR (qPCR) on a Bio-RAD
CFX Opus 95 Real-Time PCR system(Bio-Rad, Hercules, CA, USA). The gene expression
levels of total bacteria and Anammox bacteria were assessed using Axen One-step RT-qPCR
Master Mix (2X, Probe) according to the manufacturer’s instructions(Axen, Republic
of Korea). RNA was extracted from bacterial samples using the SPINeasy RNA Kit for
Bacteria(MP Biomedicals, Irvine, CA, USA) and quantified using a NanoDrop Spectrophotometer(Thermo
Scientific, Waltham, MA, USA). The extracted RNA was normalized to a concentration
of 10 ng/µL before analysis. For the qPCR assay, two primer sets were employed: one
targeting total bacteria and the other specific to Anammox bacteria, both within the
16S rRNA gene region. The primer sequences for total bacteria included Bac1055YF (5′-ATGGYTGTCGTC
AGCT-3′) and Bac1492R (5′-ACGGGCGGTGTGTAC-3′) (Harms et al., 2003), along with the probe 16STaq1115 (FAM-CAACGAGCGCAACCC-3′). For Anammox bacteria,
the primers AMX809F (5′-GCCGTAACGATGGGCACT-3′) and AMX1066R (5′-AACGCTCACGACACGAGGCTG-3′)
were used(Tsushima et al., 2007), with the probe AMX931 (FAM-TCGCACAAGGGGCTGAGCATGTGGCTT-3′) (Hamersley et al., 2007).
This table outlines the optimal thermal cycling conditions applied during qPCR reactions(Table 3). Each reaction was carried out in a total volume of 20 µL, comprising 10 µL of Axen
One-step RT-qPCR Master Mix, 1 µL of each primer (10 µM), 1 µL of probe (10 µM), and
1 µL of the template RNA. The thermal cycling protocol included an initial reverse
transcription step for cDNA synthesis at 50°C for 30 min, followed by enzyme activation
at 95°C for 10 min. Subsequently, 40 amplification cycles were carried out.
A standard curve for quantification was prepared by amplifying the target genes within
the 16S rRNA region using specific primers. The amplified gene was cloned into a plasmid
using TA cloning, and the plasmid DNA was prepared. Standard gene copy numbers were
calculated using the following equation: 1(Lee et al., 2006).
The genome size includes the plasmid and target gene sizes. The plasmid was diluted
from 1×101 to 1×107 gene copies to create a standard curve for quantifying target
genes. After running qPCR, a Ct value was obtained for each template, representing
the cycle threshold, where the fluorescent signal exceeded the background. Gene copy
numbers were calculated using the following equation: 2.
To validate the qPCR results, a standard curve was plotted using the Ct values and
the gene copy numbers. The linearity of the standard curve was evaluated by determining
Pearson’s correlation coefficient (r) and the coefficient of determination (r²)(Taylor et al., 2010).
Table 2 qPCR primer sequences
|
Taxon
|
Sequence (5’-3’)
|
Primer/Probe
|
Reference
|
|
Bacteria
|
ATGGYTGTCGTCAGCT
|
Bac1055YF
|
Harms et al., 2003
|
|
ACGGGCGGTGTGTAC
|
Bac1492R
|
|
CAACGAGCGCAACCC
|
16STaq1115 (FAM)
|
|
Anammox
|
GCCGTAAACGATGGGCACT
|
AMX809F
|
Tsushima et al., 2007
|
|
AACGTCTCACGACACGAGCTG
|
AMX1066R
|
|
TCGCACAAGCGGTGGAGCATGTGGCTT
|
AMX931 (FAM)
|
Hamersley et al., 2007
|
Table 3 Conditions of qPCR cycles used in this study
|
Genes
|
Number of cycles / Temperature / Running time
|
|
Bacteria
|
X 1 cycle
50°C
30 min
|
X 1 cycle
95°C
10 min
|
X 40 cycle
95°C 60°C
10 s 30 s
|
|
Anammox
|
3. Results and Discussion
3.1 Quantification of gene expression in Anammox bacteria using qPCR analysis
Through qPCR analysis, the transcriptional activity of Anammox bacteria was assessed
by calculating the DNA-normalized RNA expression, which is represented as the RNA/DNA
ratio. This ratio serves as a critical indicator of cellular activity, with RNA reflecting
active protein synthesis and metabolic processes, whereas DNA represents the fundamental
genetic structure of the cell. A high RNA/DNA ratio indicates active metabolism, whereas
a low ratio suggests reduced metabolic activity or dormancy. Changes in the metabolic
activity of bacteria can be observed by analyzing this ratio.
In the pre-treatment phase(Fig. 2), the C sample exhibited the highest RNA/DNA ratio of 2.22, indicating that the control
(C) retained high metabolic activity without undergoing Freeze-Thaw Cycles. In contrast,
1FT_80 and 2FT_80 samples had very low ratios of 0.159, with only 3FT_80 showing an
increase to 0.65. Similarly, 1FT_200 and 2FT_200 recorded low ratios of 0.131 and
0.175, respectively, whereas 3FT_200 exhibited a slight increase to 0.808, suggesting
some recovery in metabolic activity. Observation of the reactivation day 5 data(Fig. 2), it is evident that most samples displayed an increase in the RNA/DNA ratio after
the Freeze-Thaw Cycles compared to the pre-treatment phase. Notably, 1FT_80 showed
a substantial increase to 0.7036, whereas 1FT_200 recorded the highest ratio of 0.899,
indicating that Anammox bacteria began to recover their activity after the substrate
was introduced during the reactivation phase.
However, the 2FT_80 and 3FT_80 samples displayed a decrease in the RNA/DNA ratio,
at 0.432 and 0.252, respectively, compared to their pre-treatment values. The 2FT_200
and 3FT_200 samples also did not show significant increases, with ratios of 0.354
and 0.442, respectively. This suggests that as the number of freeze–thaw cycles increased,
some samples experienced difficulty in recovering Anammox bacterial activity. The
NFT sample, which was reactivated at 15°C without Freeze-Thaw Cycles, exhibited an
RNA/DNA ratio of 0.612. This value was lower than that of the FTC-treated samples
but still substantially lower than that of the C sample, indicating that the NFT sample
did not fully recover its metabolic activity. This comparatively low recovery may
be attributed to the absence of prior freeze–thaw stress that could have functioned
as a physiological priming stimulus. Without such preconditioning, the NFT biomass
was directly exposed to the low-temperature reactivation environment, which might
have hindered the timely activation of cold-adaptation mechanisms and reduced transcriptional
activity during the reactivation period.
Overall, the samples subjected to freeze–thaw cycles showed partial recovery of activity
on reactivation day 5, but the 2FT and 3FT samples did not exhibit the same level
of RNA/DNA increase as the 1FT samples. This indicates that Anammox bacteria faced
more challenges in recovering their activity as the number of freeze–thaw cycles increased.
In conclusion, the 1FT_200 sample showed the highest activity on reactivation day
5, whereas the NFT sample, which underwent no freeze-thaw treatment, displayed relatively
lower activity. This suggests that freeze–thaw cycles under specific conditions can
have a positive effect on the recovery of Anammox bacterial activity, but the impact
varies depending on the number of cycles.
Fig. 2. Comparison of DNA-normalized RNA expression ratios of Anammox bacteria to
total bacteria during FTC pre-treatment and on reactivation day 5. The relative RNA
expression of Anammox bacteria, normalized by DNA abundance, is calculated as the
ratio of Anammox-specific qPCR results to total bacterial qPCR results (RNA/DNA).
Data are presented for samples collected during FTC pre-treatment and after 5 days
of reactivation at 15°C.
3.2 Specific Anammox Activity results
Fig. 3 shows the SAA results after FTC on days 1 and 5 of reactivation, presented as a percentage
relative to the control. Comparing reactivation days 1 and 5, a significant increase
in SAA was observed on day 5. For example, the SAA of the 1FT_80 sample increased
from 6% on day 1 to 16% on day 5, and that of the 3FT_80 sample increased from 18%
on day 1 to 37% on day 5. Similarly, in samples frozen at -200°C, SAA increased from
5% in 1FT_200 on day 1 to 9% on day 5, and from 24% in 3FT_200 to 29% on day 5, respectively.
These results suggest that as reactivation time increased, Anammox bacteria became
more efficient at utilizing ammonium (NH₄) and nitrite (NO₂⁻) as substrates. Notably,
samples subjected to more freeze–thaw cycles initially showed lower SAA but exhibited
a gradual increase in SAA over time.
Fig. 4 presents the ammonium removal efficiency (ARE). On reactivation day 1, most samples
showed little or no ammonium removal. For instance, 1FT_80, 2FT_80, 1FT_200, and 2FT_200
samples had an ARE of 0%, whereas 3FT_80 showed a removal rate of only 3%. However,
by day 5, the situation had improved, with ARE increasing to 15% in both 1FT_80 and
2FT_80 and to 12% in 3FT_200. This indicates that the Anammox bacteria regained some
of their ability to process ammonium over time. In terms of NRE, Fig. 4 shows that there were notable differences between days 1 and 5 of reactivation. On
day 1, 3FT_80 had an NRE of 33%, which increased significantly to 74% by day 5. Similarly,
3FT_200 saw an NRE increase from 36% on day 1 to 54% on day five. In our previous
study(Kim et al., 2025), 16S rRNA-based microbial community analysis revealed that the relative abundance
of Pseudomonas significantly increased following repetitive freeze–thaw treatment.
This supports the possibility that denitrifying bacteria such as Pseudomonas were
enriched during the reactivation period. Based on this prior finding, the increase
in NRE observed in this study suggests that, in addition to Anammox bacteria, denitrifiers
such as Pseudomonas may have also contributed significantly. It is highly likely that
Pseudomonas gained a competitive advantage in nitrite substrate competition in a low-temperature
environment. Since Pseudomonas can utilize nitrite as an electron acceptor during
denitrification(Arai, 2011), the increase in NRE indicates that these denitrifying microorganisms played a significant
role during reactivation. However, in the 3FT_80 sample, the ARE decreased from 3%
on day 1 to 0% on day five. This suggests that Anammox bacteria struggled to metabolize
ammonium, possibly due to structural changes or failure of nitrite substrate competition.
Comparison of reactivation days 1 and 5, it was evident that the SAA and nitrogen
removal efficiency of Anammox bacteria generally increased over time. However, it
is important to note that an increase in SAA concentration does not necessarily indicate
a corresponding increase in the actual activity of Anammox bacteria. Anammox bacteria
metabolize ammonium (NH₄⁺) and nitrite (NO₂⁻) at a stoichiometric ratio of 1:1.32(Mulder et al., 1995; Strous, Kuenen et al., 1999). However, in the 3FT samples, while the NRE significantly increased, the ARE decreased
overall.
For instance, in the 3FT_80 sample, the NRE reached 74%, but the ARE dropped to 0%,
indicating that ammonium removal did not occur in this case. This imbalance may reflect
abnormal metabolic activity in Anammox bacteria and suggests that other microorganisms
contributed to nitrite removal during the reactivation process. In our previous study(Kim et al., 2025), 16S rRNA-based microbial community analysis showed a significant increase in the
relative abundance of Pseudomonas following repetitive freeze–thaw treatments. This
finding supports the possibility that denitrifying bacteria, such as Pseudomonas,
were selectively enriched during the reactivation period. Given that Pseudomonas can
use nitrite as an electron acceptor during denitrification(Arai, 2011), it is likely that they gained a competitive advantage in nitrite substrate utilization
under low-temperature conditions. Ultimately, Pseudomonas may have compensated for
the reduced function of Anammox bacteria, thereby maintaining high nitrogen removal
efficiency despite a decline in Anammox activity.
In conclusion, after freeze–thaw cycles, Anammox bacteria experienced some degree
of reactivation. However, due to structural changes or substrate competition for ammonium
and nitrite, their activity was limited, and denitrifying bacteria, such as Pseudomonas,
may have played a critical role in maintaining nitrogen removal efficiency.
Fig. 3. SAA relative to control on reactivation days 1 and 5. The SAA of samples treated
with FTC at -80°C and -200°C, as well as NFT samples (reactivated at 15°C without
FTC), is expressed as a percentage relative to the control (C). SAA measurements were
taken on reactivation days 1 and 5 to assess changes in Anammox bacterial activity.
Fig. 4. Ammonium and Nitrite Removal Efficiencies (ARE and NRE) on reactivation days
1 and 5. The Ammonium Removal Efficiency (ARE, %) and Nitrite Removal Efficiency (NRE,
%) for samples treated with FTC at -80°C and -200°C, as well as NFT samples (reactivated
at 15°C without FTC). Data are shown for reactivation on days 1 and 5.
4. Conclusion
This study evaluated the impact of repetitive FTC on the acclimation capacity of Anammox
bacteria during low-temperature reactivation. The assessment focused on the survival
and functional recovery of bacteria, primarily through the analysis of their metabolic
activity. qPCR analysis revealed a decline in the RNA/DNA ratio, reflecting reduced
transcriptional activity, metabolic potential, and survival of Anammox bacteria. The
results demonstrated that under low-temperature reactivation conditions in the presence
of substrates (ammonium and nitrite), the metabolic activity and growth of FTC-pretreated
Anammox bacteria were significantly constrained. In addition, SAA increased with the
number of FTC cycles compared to the Control (30°C) and NFT (15°C without FTC), but
the increase was primarily driven by NRE, while ARE remained consistently low. This
discrepancy further supports the interpretation that the observed metabolic activity
did not fully reflect functional recovery of Anammox bacteria, particularly with regard
to ammonium oxidation.
In a previous study, FTC pre-treatment without substrate (ammonia and nitrite) effectively
enriched Anammox bacteria in low-temperature environments(Park et al., 2024). In contrast, this study found that during low-temperature reactivation with substrate
addition, the metabolic activity and growth of FTC-treated Anammox bacteria were limited.
The initial hypothesis anticipated that FTC would enhance Anammox activity during
the reactivation phase; however, the experimental results demonstrated a decline in
performance with increasing FTC cycles. This outcome—contrasting with the original
hypothesis—highlights the physiological vulnerability of Anammox bacteria to repetitive
low-temperature stress. Furthermore, FTC appears to act not as a stimulatory factor
but rather as a selective pressure shaping microbial competition and survival.
In summary, although FTC pre-treatment contributed to the initial selective enrichment
of Anammox bacteria under low-temperature conditions, it was insufficient to ensure
sustained metabolic activity and functional recovery during the reactivation phase.
The observed increase in SAA was primarily attributed to nitrite removal, whereas
consistently low ammonia removal efficiency and a decline in the RNA/DNA ratio indicated
a reduction in the metabolic potential and adaptive failure of Anammox bacteria. These
findings suggest that stable operation of the Anammox process under cold conditions
requires more than simple pre-treatment strategies; rather, a comprehensive approach
that considers microbial interactions and responses to environmental stressors is
essential for enhancing process resilience and reactivation efficiency. These insights
deepen our understanding of microbial community dynamics and provide practical guidance
for designing low-temperature Anammox processes with improved resilience.
Acknowledgement
This work is financially supported by Korea Ministry of Environment(MOE) Graduate
School specialized in Integrated Pollution Prevention and Control Project.
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