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. 2026 Feb 13;12:101349. doi: 10.1016/j.crfs.2026.101349

Impact of kilning on the fermentability of oat wholemeal flour and related bioconversion of starch, protein, dietary fibre, and phytate

Eline Lambrechts a,1, Ewoud Blontrock a,1, Thomas Gettemans b, Jolien Lemoine a, Sarah Vanhove a, Yamina De Bondt a, Frederik Janssen a, Stefan Weckx b, Luc De Vuyst b, Arno GB Wouters a, Christophe M Courtin a,⁎
PMCID: PMC12927064  PMID: 41737888

Abstract

Considering the nutritional profile of whole oats and the potential health-enhancing effect of fermentation, oat fermentation offers significant potential for developing nutritious foods. However, fermenting oat wholemeal flour, in particular with lactic acid bacteria, proves challenging. We hypothesise this to be related to the kilning step, a heat-treatment typically applied to intact oat groats, which inactivates oat endogenous enzymes, enhancing stability against lipid oxidation and rancidity. However, kilning may also limit enzyme-induced biochemical changes of the food matrix and the enzymatic release of fermentable substrates during fermentation, thereby potentially impeding microbial growth and acidification. To investigate this, oat wholemeal flour suspensions derived from kilned and non-kilned oat groats were fermented for maximum 24h using a strain of Lactiplantibacillus plantarum as a starter culture. The ferments were comparatively evaluated microbiologically (i.e., microbial growth and starter culture prevalence) and biochemically (i.e., acidification, metabolite content, soluble fibre content, soluble protein content, and phytate reduction) over time. Fermentation proceeded faster in non-kilned oat wholemeal flour suspensions, resulting in more rapid acidification and more pronounced biochemical changes than in kilned suspensions. These results highlight the importance of active oat endogenous enzymes to provide nutrients for the fermenting lactic acid bacteria and to achieve desired biochemical changes of the matrix during oat fermentation.

Keywords: Oat, Kilning, Lactic acid fermentation, Starch, Protein, Fibre, Phytate

Graphical abstract

Image 1

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Highlights

  • •

    Lactiplantibacillus plantarum Vega Boost LP acidifies and grows much faster in non-kilned than in kilned oat wholemeal flour suspensions.

  • •

    Kilning reduces oat enzyme activities, limiting molecular changes in oat constituents during fermentation.

  • •

    Activity of oat enzymes is crucial for generating nutrients for the fermenting bacteria.

Abbreviations

K-DOW = 

kilned defatted oat wholemeal flour

NK-DOW = 

non-kilned defatted oat wholemeal flour

CFU = 

Colony-forming units

AX = 

arabinoxylan

P = 

phosphate

1. Introduction

Cereal-based fermented food products such as sourdough bread and plant-based dairy alternatives are gaining more attention as fermentation is considered a potential tool to enhance the nutritional profile of cereal products (Arora et al., 2021; Dhakal et al., 2023; Poutanen et al., 2009; Ribet et al., 2023). Previous research has indicated that lactic acid fermentation of cereal-based raw materials (e.g. wholemeal wheat and rye flour) results in dietary fibre solubilisation (Coda et al., 2014; Lappi et al., 2010a, Lappi et al., 2010b; Lu et al., 2019; Mihhalevski et al., 2013), phytate degradation (Fekri et al., 2020; Leenhardt et al., 2005; Mariotti et al., 2014; Reale et al., 2007), and protein hydrolysis (Loponen et al., 2004; Thiele et al., 2004; Tuukkanen et al., 2005) which are linked with enhanced gut health, improved mineral bioaccessibility, and increased protein digestibility, respectively, of derived food products such as sourdough bread. The responsible biochemical reactions are mostly catalysed by cereal endogenous enzymes such as endoxylanases, β-glucanases, phytases, and proteases, all of which show pH-dependent activities (Fu et al., 2024; Nkhata et al., 2018). This is relevant in the context of lactic acid fermentation, whereby the pH typically decreases from around neutral pH to below pH 4.5. For example, Leenhardt et al. (2005) showed that endogenous wheat phytase is predominant over microbial phytase in reducing phytic acid content during wheat sourdough breadmaking. Arte et al. (2015) reported a similar degree of protein hydrolysis during both fermentation and incubation at pH 4.5 in the presence of antibiotics of wheat bran, indicating that endogenous proteases are mostly responsible for protein hydrolysis. Besides the potential nutritional benefits, these enzymatic conversions are also crucial to provide sufficient nutrients for the fermenting lactic acid bacteria (LAB). Most cereal substrates are low in fermentable saccharides and require the additional hydrolysis of starch by endogenous amylases (Fu et al., 2024; Gobbetti and Gänzle, 2023). Additionally, LAB need an adequate supply of amino acids to support their growth, withstand highly acidic environments, and maintain stable homeostasis in their stationary growth phase (Gänzle, 2014). In the context of wheat and rye fermentations, it has been suggested that LAB rely on the primary hydrolysis of complex proteins by cereal proteases into smaller oligopeptides, which can be further metabolised by microbial intracellular peptidases (Arte et al., 2015; Gänzle et al., 2008). Considering the above, raw materials with low endogenous enzyme activities are expected to have a poor fermentation quality.

Amongst cereals, oats are a promising raw material for lactic acid fermentation because of their beneficial nutritional profile. Oats have a high protein content, a relatively well-balanced amino acid profile, and are naturally gluten-free (Alemayehu et al., 2023; Tang et al., 2023). Additionally, oats are rich in β-glucans to which cholesterol-lowering and blood glucose-lowering effects are ascribed (EFSA, 2011). Previous research already indicated that fermented oat products (e.g. fermented oat drink or oat-based yoghurt) can have several nutritional improvements, including increased probiotic content and improved antioxidant and anti-inflammatory properties (Alemayehu et al., 2023; Djorgbenoo et al., 2023; Yu et al., 2023). Nevertheless, few fermented oat products are currently available in European supermarkets (Djorgbenoo et al., 2023). Furthermore, most of these products are made by fermenting a liquid oat base, in which case fermentation is carried out after prior extraction of soluble constituents and thus after removing the insoluble, yet nutritionally valuable constituents. As a result, little is known about the impact of fermentation on major oat constituents such as starch, protein, fibre, and phytate.

Almost all oat kernels are industrially heat-treated (i.e., kilned) to inactive lipase and, as such, prevent rancidity development during processing and storage (Decker et al., 2014). This heat-treatment also inactivates other oat endogenous enzymes and could therefore impact the bioconversion reactions of starch, protein, and fibre during fermentation. Recently, it was demonstrated that the biochemical changes induced by oat endogenous enzymes at pH values relevant for lactic acid fermentation are very limited in oat wholemeal flour derived from kilned oat groats (Blontrock et al., 2025). Other studies have already reported on the poor fermentability of oat-based raw materials. For example, Loponen et al. (2007) did not observe any protein hydrolysis during fermentation of oat bran. Wolter et al. (2014) showed that Weissella cibaria was unable to grow in oat flour during fermentation, which was related to the low availability of fermentable saccharides due to the lack of endogenous α-amylase activity (Wolter et al., 2014). Both studies have explained these results by the prior heat-treatment of oat groats, resulting in a lack of oat endogenous protease and amylase, respectively (Loponen et al., 2007; Wolter et al., 2014).

Reaching a pH of 4.5 or below in a short timeframe is a prerequisite for preventing outgrowth of pathogens which might be present in the raw material. The fermentability of oat wholemeal flour has previously been improved by adding oat or barley bran, as a source of minerals (Grgić et al., 2024), additional fermentable saccharides (Alharbi et al., 2022), or exogenous α-amylases (Luana et al., 2014). To the best of our knowledge, the use of non-kilned oats, which still contain all endogenous enzyme activities, has not yet been explored in this regard. Therefore, the goal of this study was to systematically compare the fermentability (i.e., acidification, microbial growth, starter culture prevalence and metabolite production) of oat wholemeal flour derived from both kilned and non-kilned oat groats and study the associated changes in oat starch, protein, fibre, and phytate. The main hypothesis is that oat kilning strongly reduces the fermentability of oat wholemeal flour and therefore hampers biochemical changes in oat starch, protein, fibre, and phytate.

2. Materials & methods

2.1. Materials

The same kilned and non-kilned oat wholemeal flours were used as those described in Blontrock et al. (2025) and defatted, characterised for their constituent composition, and endogenous enzyme activities as described therein. Defatting did not have an impact on acidification and microbial growth during wholemeal flour fermentation (as can be seen in Figure S.1 in the supplementary data).

A commercial homofermentative strain of L. plantarum (Vega Boost LP) was used as a starter culture for fermentation and was kindly provided by Chr. Hansen (part of Novonesis, Hoersholm, Denmark).

All chemicals, solvents, and enzymes used in this study were of analytical grade and were obtained from Chem-lab Analytical (Zedelgem, Belgium), VWR International (Leuven, Belgium), Fisher Scientific (Brussels, Belgium), Megazyme (Wicklow, Ireland), and Merck (Bornem, Belgium).

2.2. Methods

2.2.1. Fermentation of oat wholemeal flour

Preparation of inoculum. A single colony obtained after plating of the stater culture on de Man, Rogosa, and Sharp (MRS) agar medium was inoculated in 10.0 mL of liquid MRS growth medium for 24 h at 30 °C while shaking (250 rpm) to reach a maximal cell density. The obtained liquid culture was sub-cultured at 1.0% (v/v) for 10 h under the same conditions in 10.0 mL of fresh MRS medium. The obtained bacterial suspension was centrifuged at 4000×g for 7 min, and the pellet washed with a saline solution (0.85% w/v), and centrifuged before re-suspending in saline solution. This suspension was used for inoculation of the oat flour-water suspension.

Oat fermentations. Two oat-based raw materials, kilned defatted oat wholemeal flour (K-DOW) and non-kilned defatted oat wholemeal flour (NK-DOW), were weighed (7.0 g) into sterilised screwcap bottles. Next, 41.0 mL of sterile ultrapure water (Milipore, Burlington, MA, USA) and 490 μL of the inoculum (to reach an initial cell density of 107 CFU/mL suspension) were added. This resulted in a flour suspension at a 1:6 (w/v) flour-to-water ratio, which was necessary to obtain homogenous mixing. Fermentation processes were conducted for 24 h in a water bath at 30 °C under continuous magnetic stirring (300 rpm). The incubation temperature was selected based on the strain's optimal growth temperature, following information provided by the supplier. Independent and individual fermentations were included for every timepoint, in triplicate, with every replicate performed on a different day.

Sampling. Samples were taken before inoculation (0 h), immediately after inoculation (0’ h), and after 2, 4, 8, 12, and 24 h of fermentation. At each time point, one bottle was removed from the water bath for each ongoing fermentation. The suspension was further subdivided into five parts by weighing:

  • i.

    1.0 g sample for selective plating.

  • ii.

    5.0 g sample for culture-independent microbiological analysis. The sample was centrifuged at 1000×g for 5 min to remove debris originating from the DOW. The obtained supernatant was split into 2 equal fractions and further centrifuged at 5000×g for 20 min to obtain a cell pellet, which was subsequently stored at −20 °C until further DNA extraction.

  • iii.

    5.0 g sample for metabolite analysis. The sample was diluted 1:5 with ultrapure water and centrifuged at 5000×g for 20 min. The supernatant was stored at −20 °C until further analysis.

  • iv.

    20.0 g sample for physicochemical characterisation. The sample was diluted 1:0.6 with ultrapure water (to reach 1:10 w/v) and centrifuged for 10 min at 10,000×g. The supernatant was heat-treated (100 °C, 10 min) to inactivate residual enzymatic activity completely.

  • v.

    The remaining suspension was frozen in liquid nitrogen and freeze-dried to obtain a fermented DOW powder.

An overview of the sampling scheme used in this study is shown in Fig. 1.

Fig. 1.

Fig. 1

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Overview of the sampling methodology during fermentation and the subsequently conducted analyses. DOW = defatted oat wholemeal flour, P = phosphate, K = kilned, NK = non-kilned.

Acidification and microbial growth profiles. The pH was continuously monitored over the course of the 24 h fermentations by an iCinac LAB fermentation monitor system (KPM analytics, Westborough, USA). Additionally, at each sampling point, the microbial growth of LAB and yeasts during the fermentation was monitored using selective plating of appropriate serial dilutions of the fermented DOW suspension (Sample i, Fig. 1) on MRS-5 agar medium (Meroth et al., 2003) supplemented with 100 ppm of cycloheximide (Sigma-Aldrich, St Louis, USA) and 5 ppm of amphotericin B (Sigma-Aldrich), and Yeast extract-Peptone-Glucose (YPG) agar medium supplemented with 100 ppm of chloramphenicol (Sigma-Aldrich), respectively. Plates were incubated for 48 h at 30 °C and colonies were counted, and recalculated to colony-forming units (CFU)/g of DOW suspension.

2.2.2. Culture-independent microbiological analysis

To determine whether the inoculated starter culture became the most dominant microorganism during the oat-based fermentations the microbial composition of the ferments was analysed. Given that this is an inoculated fermentation with a relatively short duration (24h), other in-depth analyses (e.g. α- and β-diversity) were considered unnecessary for the current study. The DNA extraction of the cell pellet (Sample ii, Fig. 1) was performed as described previously in Pradal et al. (2024). In short, one cell pellet per timepoint was thawed and subjected to a combination of enzymatic lysis using mutanolysin (Sigma-Aldrich), lysozyme (Merck, Darmstadt, Germany) and mechanical lysis steps. Further purification of the extracted DNA was done using the DNAeasy Blood & Tissue kit (Qiagen, Hilden, Germany) and DNA concentrations were measured using a Qubit fluorimetry system (1X dsDNA High Sensitivity kit, Thermo Fisher Scientific). PCR amplification of the full-length 16S rRNA gene was performed as described previously by Decadt et al. (2023) using barcoded 27F (5′-AGRGTTYGATYMTGGCTCAG-3′) and 1492R (5′-RGYTACCTTGTTACGACTT-3′) primers (Integrated DNA Technologies, Coralville, IA, USA). The amplification success was evaluated using agarose gel electrophoresis on a 1.5-% (m/v) agarose gel at a constant voltage of 3.33 V/cm for 1 h. PCR mixtures were purified using the Wizard Plus SV Minipreps DNA purification system (Promega, Madison, WI, USA) and concentrations were measured using Qubit fluorimetry. Sequencing was performed after equimolar pooling of the samples on a PacBio Sequel IIe system (VIB Nucleomics Core Facility, Leuven, Belgium). The sequence reads were processed using RStudio (version 4.2.1; RStudio Team, 2022) and amplicon sequence variants (ASVs) were obtained using the DADA2 package (version 1.26.0, Callahan et al., 2017). The used filtering parameters were minQ = 3, minLen = 1100, maxLen = 1600, maxN = 0, and maxEE = 2. Taxonomy assignment was based on the SILVA database (version 138.1; Quast et al., 2013) using a minBoot = 80, after which mitochondrial and chloroplast reads were filtered out. ASVs were either grouped by species for general representation of the bacterial species composition or represented as such in case of the starter culture species for starter strain follow-up throughout the fermentation. Rarefaction analysis was performed on the species level using the R package vegan (v2.6-4, Oksanen et al., 2025). For visual representation of the species composition, species with a relative abundance below 5 % in all samples were grouped together under the category ‘Minorities <5 %’.

2.2.3. Analysis of free saccharide dynamics

To study the production of fermentable saccharides and their subsequent consumption by the fermenting microorganisms, the mono-, di-, and oligosaccharide composition of the fermented DOW suspensions was determined similarly as in Langenaeken et al. (2019). For this, heat-treated fermented supernatants (Sample iv, Fig. 1) were diluted 100 times in ultrapure water, filtered through a Millex-GP 0.22 μM PES syringe filter (MilliporeSigma, Burlington, MA, USA) and transferred into vials. Glucose, fructose, sucrose, maltose, and maltotriose contents were analysed with a Dionex ICS-5000 High-Performance Anion Exchange Chromatography system with Pulsed Amperometric detection (HPAEC-PAD, Sunnyvale, CA, USA), using a PA-100 analytical column (4 × 250 mm). The elution solvent consisted of 100 mM NaOH at a flow rate of 1.0 mL/min for 5 min with a gradual increase of sodium acetate at 3.6 mM/min for the next 25 min of the run. Quantification was performed by adding rhamnose as an internal standard and by including a calibration standard solution containing known concentrations of rhamnose, glucose, fructose, sucrose, maltose, and maltotriose.

2.2.4. Analysis of metabolite production

Concentrations of chemical compounds produced during the fermentation were quantified in Sample iii (Fig. 1). A suite of chromatographic techniques was employed for the quantification of major fermentation-related metabolites. Sugar alcohols were quantified through HPAEC-PAD using a CarboPac MA1 column on an ICS 5000 chromatograph equipped with an ED-40 PAD detector (Dionex). Sample preparation and run conditions were as described by Pradal et al. (2024). Organic acid concentrations were determined through ultra-performance liquid chromatography with tandem mass spectrometry detection (UPLC-MS/MS) using an HSS T3 column on a Waters Aquity UPLC system with a tandem mass spectrometer (Waters, Milford, MA, USA) as described by González-Alonso et al. (2024), with the modification that lactic acid was also determined using this technique. Concentrations of acetic acid and ethanol were determined using gas chromatography with flame ionisation detection (GC-FID) with a Trace Finder 1310 gas chromatograph equipped with a DB wax UI column and an FID-80 detector (Thermo Fisher Scientific, Waltham, MA, USA) as described by González-Alonso et al. (2024). All samples were run in triplicate with external calibration using pure compounds.

2.2.5. Analysis of protein extractability and apparent molecular weight distribution

The protein content (N x 5.83) in heat-treated fermented supernatants (Sample iv, Fig. 1) was quantified using a total combustion method (Dumas elemental analyser, EA1108 model, Carlo Erba, Hindley Green, UK). Protein extractabilities were calculated as described in Blontrock et al. (2025). Size-exclusion high-performance liquid chromatography (SE-HPLC) was used to evaluate the protein molecular weight distribution before and after fermentation, as described by Blontrock et al. (2025). In short, freeze-dried fermented DOW suspensions were dispersed, and heat-treated extracts were diluted in a sodium phosphate buffer with 2.0% w/v sodium dodecyl sulfate and 1.0% w/v 1,4-dithiothreitol, filtered (0.45 μm), and injected onto a Biosep SEC-S2000 column mounted on a Shimadzu (Kyoto, Japan) modular chromatography system for separation. Chromatograms were recorded using a UV-Vis detector at 214 nm, and results were normalised for injection volume and the amount of starting material for the wholemeal flour suspensions, allowing comparison between sample types. As protein molecular weight markers, bovine serum albumin (66.5 kDa), carbonic anhydrase from bovine erythrocytes (30 kDa), α-lactalbumin (14 kDa), aprotinin (6.5 kDa), and (Ala)5 (373 Da) were analysed under the same experimental conditions as the samples. Expected elution times are 10.0 – 12.0 min for oat globulin monomers, 12.5 min for the oat globulin α-subunit, 13.0 min for the oat globulin β-subunit, 14.0 min for albumins/avenins/degraded oat globulins, and longer than 15.0 min for smaller peptides (Blontrock et al., 2025).

2.2.6. Analysis of arabinoxylan extractability

The arabinoxylan (AX) content of the heat-treated fermented supernatants (Sample iv, Fig. 1) was determined following the procedure described by Courtin et al. (2000). Samples were diluted in 4.0 N trifluoracetic acid to fully hydrolyse carbohydrates into monosaccharides. Next, the monosaccharides were reduced to alditols and derivatised into alditol acetate using NaBH4 and acetic acid anhydride, respectively. Finally, the alditol acetates were analysed using GC-FID (6890N, Agilent Technologies, Santa Clara, CA, USA) with the aid of a Supelco SP-2380 separation polar column (30 m × 0.32 mm, 0.2 μm film thickness, Bellefonte, PA, USA). Concentrations of xylose and arabinose were determined using allose as an internal standard and an external calibration solution with known concentrations of arabinose and xylose. Both these concentrations were summed and multiplied by 0.88 (correction for water hydrolysis) to obtain the total AX concentration in the supernatants. AX extractability was calculated by dividing the amount of AX recovered in the heat-treated fermented supernatants by the initial total AX content of the DOW sample, weighed at the start of fermentation.

2.2.7. Analysis of β-glucan extractability

The β-glucan content in DOW and heat-treated fermented supernatants (Sample iv, Fig. 1) was quantified using the mixed-linkage β-glucan kit (Megazyme, Wicklow, Ireland; AACC Method 32-23.01) with some minor modifications, as explained by Blontrock et al. (2025). Extracted β-glucan was precipitated by diluting an aliquot of the heat-treated fermented supernatants in ethanol to reach a final ethanol concentration of 90% v/v, followed by centrifugation (10 min at 1800×g), after which the resulting pellet was resuspended in sodium phosphate buffer (0.020 M, pH 6.5). Hydrolysis steps with lichenase and β-glucosidase were then performed, followed by glucose quantification using glucose oxidase/peroxidase reagent and spectrophotometric analysis (510 nm, Ultraspec, 2000 UV/VIS). To control for free glucose in the samples, a blank (prepared by treating the sample with lichenase but not β-glucosidase before reaction with the glucose oxidase/peroxidase reagent) was included. A glucose standard solution (1.5 mg/mL) was used to correlate absorbance with glucose concentration. β-glucan extractability was defined as explained for arabinoxylan (section 2.2.6).

2.2.8. Analysis of phytate degradation

The degree of phytate degradation was determined as the ratio of free over total (free and myo-inositol-bound) phosphate, as previously explained by Blontrock et al. (2025). For this, 500 mg of Sample v (Fig. 1) was diluted in 0.66 M HCl (1:10 w/v) and shaken for at least 16 h. Next, samples were centrifuged (13000 rpm, 10 min), and the obtained supernatant was diluted (1:1 v/v) in 0.75 M NaOH. To determine the total phosphate content, phytic acid was completely hydrolysed into myo-inositol and phosphate using phytase and alkaline phosphatase. The total amount of phosphate was quantified after the reaction with an ammonium molybdate colour reagent (including ascorbic and sulphuric acid) and absorbance measurement at 655 nm using an Ultraspec 2000 UV/VIS spectrophotometer. To determine the free phosphate content, the same procedure was followed but all enzyme solutions were replaced with ultrapure water.

3. Results

3.1. Acidification and microbial growth during fermentation

Similar pH profiles and microbial growth curves were obtained for the independent replicates of the different oat-based suspension fermentations (Fig. 2). The pH decreased from an initial value of 6.3 ± 0.1 to 4.3 ± 0.1 after 24 h in K-DOW suspensions and from pH 6.3 ± 0.1 to pH 3.4 ± 0.1 in NK-DOW suspensions. The acidification was considerably faster during the fermentation of NK-DOW suspensions, reaching pH values below 4.5 within the first 6 h of fermentation. In contrast, fermentation of K-DOW suspensions only reached pH values below 4.5 after 14 h of fermentation (Fig. 2A).

Fig. 2.

Fig. 2

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Acidification profile (A) and bacterial cell counts (on MRS-5) in log units (B) of defatted oat wholemeal flour suspensions prepared from kilned (K-DOW) and non-kilned (NK-DOW) oat groats during a 24 h fermentation with Lactiplantibacillus plantarum Vega Boost LP. Replicates on three separate days are indicated by the letters A, B, and C.

The start inoculum cell count was at 7.4 ± 0.1 log CFU/g suspension in both K-DOW and NK-DOW suspensions (slightly above the targeted 7.0 log CFU/g) and increased to 8.9 ± 0.1 log CFU/g suspension in K-DOW and to 9.5 ± 0.1 log CFU/g suspension in NK-DOW (Fig. 2B). Until time point 4 h, the bacterial counts increased similarly in both K-DOW and NK-DOW suspensions. Thereafter, bacterial growth continued exponentially until 12 h of fermentation in NK-DOW suspensions, whereas the microorganisms in the K-DOW suspensions started transitioning into the stationary phase around 8 h into the fermentation. This indicates that the bacteria grew less well in K-DOW compared to NK-DOW suspensions. Yeasts were not found in the different fermentations at any of the time points except for some very low counts in some samples (data not shown), effectively indicating a very low presence of yeasts overall.

3.2. Starter culture prevalence during fermentation

Rarefaction analysis showed, for those samples where sequencing data was available, that all samples were sequenced with sufficient sequencing depth to reliably assess the bacterial composition of the samples (Figure S.2 in the supplementary data).

The bacterial species diversity before inoculation (time point 0 h) of the NK-DOW suspensions, though based on a single observation due to technical limitations while amplifying the microbial DNA, shows a very diverse range of different microorganisms, including background L. plantarum strains (Fig. 3A). The mentioned limitations mainly originate from the presence of unknown quantities of non-bacterial DNA after DNA extraction, which is more pronounced in samples with a low bacterial load. As the employed DNA extraction protocol ensures lysis of all bacterial cells, other cells can be lysed as well, leading to extracted DNA that does not yield adequate concentrations of 16S rRNA gene amplicons. Due to this, pre-inoculation microbial compositions for K-DOW suspensions could also not be retrieved. However, the diversity originating from the NK-DOW suspensions is speculated to be higher than the initial diversity of the K-DOW suspension, as the latter went through a kilning step. After inoculation, the relative abundance of L. plantarum increased from a background species to the most abundant species in the NK-DOW suspension. Inoculated K-DOW suspensions did not allow the retrieval of enough PCR amplicons for sequencing. During fermentation, the bacterial diversity was reduced to a major presence of L. plantarum, with more than 94% of all reads being attributed to this species in all samples after 8 h.

Fig. 3.

Fig. 3

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Culture-independent microbial analysis of the bacteria in defatted oat wholemeal flour suspensions prepared from kilned (K-DOW) and non-kilned (NK-DOW) oat groats during a 24 h fermentation with Lactiplantibacillus plantarum Vega Boost LP, based on the full-length 16S rRNA gene. Replicates on three separate days are indicated by the letters A, B, and C. Only sampling points 0 h, 0′ h, 8 h, and 24 h were analysed. Asterisks (∗) represent samples for which no sufficient amplicon concentration could be obtained for sequencing. Overview of the bacterial species present in the samples with the reads obtained per sample during sequencing shown as a bar plot on top of the relative abundance plot (A). ASVs of the starter culture species, Lactiplantibacillus plantarum, throughout the different fermentations (B).

At the ASV level, five different ASVs of the 16S rRNA gene were found in equal ratios within the reads of L. plantarum before inoculation in NK-DOW suspensions, reflecting the average of five copies of the gene in the species. The ASVs found in all sequenced samples after inoculation were not distinct from the background L. plantarum, which prevents discriminating the starter culture strain from the background strains and thus, the absolute conclusion that the starter culture strain prevailed. However, the combination of the sharp increase in bacterial counts after inoculation and L. plantarum being the dominant species throughout all replicates of the fermentations with K-DOW and NK-DOW suspensions, makes it reasonable to assume that the inoculated strain was indeed dominant during fermentation of both K-DOW and NK-DOW suspensions.

3.3. Production and consumption of fermentable saccharides during fermentation

Initially, K-DOW and NK-DOW suspensions contained a similar amount of fermentable saccharides, of which most was sucrose (1.3 ± 0.1% dm m/mDOW). K-DOW suspensions also contained low amounts of glucose (0.03 ± 0.01%), fructose (0.03 ± 0.01%) and maltose (0.01 ± 0.01%) while NK-DOW suspensions contained slightly higher amounts of glucose (0.05 ± 0.01%), fructose (0.04 ± 0.01%) and maltose (0.05 ± 0.01%). The amount and type of fermentable saccharides derived from starch or other macromolecules during the fermentation of oat flour suspensions differed strongly between K-DOW and NK-DOW (Fig. 4). K-DOW suspensions showed a gradual decrease in sucrose content over time, but other fermentable saccharides were not detected after 2 h. In contrast, NK-DOW suspensions showed a faster decrease in sucrose over time and an increase in glucose, fructose, maltose and maltotriose during the first 4 h of fermentation, followed by a decrease until 24 h. Interestingly, fermentable saccharides were completely depleted after 12 h of fermentation in NK-DOW suspensions but not yet in K-DOW suspensions, suggesting the rate of fermentable saccharide consumption was faster in NK-DOW compared to in K-DOW fermentations (Fig. 4).

Fig. 4.

Fig. 4

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Glucose, fructose, sucrose, maltose and maltotriose contents in a defatted oat wholemeal flour suspension prepared from kilned (K-DOW) and non-kilned oat groats (NK-DOW) during a 24 h fermentation with Lactiplantibacillus plantarum Vega Boost LP. Dots and error bars represent the average and standard deviation of three individual and independent fermentation replicates. dm = dry matter.

3.4. Metabolite production during fermentation

Overall, the metabolite concentrations were low over the course of fermentation except for lactic acid concentrations, which increased up to 2.56 ± 0.08% dm after 24 h of fermentation for K-DOW and up to 8.02 ± 0.19% dm for NK-DOW. The rate of lactic acid production was faster for NK-DOW than for K-DOW fermentation (Fig. 5A). Concentrations of xylitol, mannitol, erythritol, sorbitol, ethanol, acetic acid, succinic acid, fumaric acid, and glucuronic acid were below the limit of quantification in all samples, as expected in a fermentation process involving solely the homofermentative metabolism of L. plantarum. Arabitol and citric acid were present at very low concentrations (below 0.0014% dm), and only limited variations over time were observed. Both myo-inositol and glycerol increased during the fermentations of NK-DOW suspensions but not in the K-DOW suspensions, indicating the production of these compounds, either by endogenous enzymes or by the starter culture strain (Fig. 5B and C). Conversely, the small amount of malic acid present in the suspensions was depleted after 4 h and 12 h for NK-DOW and K-DOW suspensions, respectively (Fig. 5D).

Fig. 5.

Fig. 5

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Lactic acid (A), myo-inositol (B), glycerol (C) and malic acid (D) contents of defatted oat wholemeal flour suspensions prepared from kilned (K-DOW) and non-kilned (NK-DOW) oat groats during a 24 h fermentation with Lactiplantibacillus plantarum Vega Boost LP. Dots and error bars represent the average and standard deviation of three individual and independent fermentation replicates. Samples with missing values are samples where the concentrations were below the limit of quantification. dm = dry matter.

3.5. Molecular changes in fibre, protein and phytate during fermentation

β-glucan. The extractability of β-glucan initially increased rapidly during the first hours of the NK-DOW fermentation, from an initial value of 29.8 ± 1.6% to 70.4 ± 1.3% at 2 h (Fig. 6A). However, as fermentation continued, the extractability of β-glucan seemingly gradually decreased, reaching 44.5 ± 0.4% after 24 h. In contrast, K-DOW suspensions exhibited a steady increase in β-glucan extractability throughout the 24-h fermentation period, starting at 13.6 ± 0.5% and rising to 43.4 ± 1.6%.

Fig. 6.

Fig. 6

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(A) β-glucan solubilisation expressed as β-glucan extractability over time, (B) arabinoxylan solubilisation expressed as arabinoxylan extractability over time, (C) phytate degradation expressed as free/total phosphate content over time and (D) protein solubilisation expressed as protein extractability over time of defatted oat wholemeal flour suspensions prepared from kilned (K-DOW) and non-kilned (NK-DOW) oat groats during a 24 h fermentation with Lactiplantibacillus plantarum Vega Boost LP. Extractabilities are defined as the percentage of constituent recovered in the extracts relative to the constituent content in an equivalent amount of defatted oat wholemeal flour (DOW). Dots and error bars represent the average and standard deviation of three individual and independent fermentation replicates. For K-DOW suspensions, only sample points 0 h, 8 h, and 24 h were analysed since the differences between 0 h and 24 h were very small.

Arabinoxylan. AX extractability increased strongly over time during fermentation of NK-DOW suspensions, rising from 9.4 ± 0.2% at 0 h to 29.0 ± 1.1% after 24 h (Fig. 6B). In K-DOW suspensions, a limited increase only from 8.4 ± 0.6% at 0 h to 14.4 ± 0.3% at 24 h was observed.

Phytate. During fermentation of NK-DOW suspensions, the ratio of free over total phosphate increased linearly from 9.1 ± 0.3% at 0 h to 35.0 ± 4.4% after 24 h, indicating a gradual hydrolysis of phytate over time (Fig. 6C). For K-DOW suspensions, a slight decrease over time was observed.

Protein. Protein extractability remained essentially constant at around 17% for the first 8 h during fermentation of NK-DOW suspensions (Fig. 6D). In the following period, a slight increase was observed, reaching a protein extractability of 25.8 ± 0.5% by the end of the 24-h fermentation. Conversely, the fermentation of K-DOW suspensions did not result in any substantial change in protein extractability, remaining at 4.2 ± 0.1% after 24 h. As shown in Fig. 7 for both non-fermented and fermented K-DOW suspensions, the differences in protein molecular weight distributions were limited. Despite the presence of some protein hydrolysis products, eluting after 21.5 min, changes in protein molecular weight distributions for NK-DOW during 24h fermentation were also limited.

Fig. 7.

Fig. 7

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Representative size-exclusion high-performance liquid chromatography profiles of proteins in defatted oat wholemeal flour suspensions prepared from kilned (K-DOW) and non-kilned (NK-DOW) oat groats during fermentation with Lactiplantibacillus plantarum Vega Boost LP, analysed under denaturing and reducing conditions at 1.0 mg protein/mL. To remove the elution peak of 1,4-dithiothreitol (occurring at 24-30 min), chromatograms were truncated at 24 min. Molecular weight markers (from left to right: 66.5 kDa, 30.0 kDa, 14.0 kDa, 6.5 kDa, 373 Da) are indicated at their corresponding elution times. AU = absorbance units.

4. Discussion

4.1. Impact of kilning on the fermentability of oat wholemeal flour

The fermentation behaviour of K-DOW and NK-DOW suspensions was investigated through microbial and chemical analyses during 24 h of fermentation with L. plantarum Vega Boost LP. Analyses included pH evolution, microbial proliferation and community composition, lactic acid biosynthesis, and sugar metabolism.

NK-DOW suspensions exhibited a markedly stronger microbial growth in comparison to K-DOW (Fig. 2B). In addition, L. plantarum reached the stationary phase in K-DOW at a lower microbial density than in NK-DOW suspensions, suggesting a lower availability of carbon and/or nitrogen sources in the former case. Indeed, LAB require sufficient carbohydrates and amino acids to grow and acidify (Hayek et al., 2013). In K-DOW suspensions, only sucrose was present and slowly depleted over time, while in NK-DOW suspensions, glucose, fructose, maltose, and maltotriose were also present, due to the partial degradation of starch by endogenous amylases (Fig. 4). In addition, the faster depletion of all fermentable saccharides in the NK-DOW suspensions (after 12 h) compared to K-DOW (after 24 h) might indicate a more efficient fermentation metabolism in the former. The protein apparent molecular weight profiles showed a greater amount of low molecular weight proteins present in fermented NK-DOW after 24 h, suggesting a higher availability of small peptides and amino acids for the fermenting LAB compared to K-DOW (Fig. 7). In NK-DOW suspensions, oat endogenous proteases can hydrolyse oat proteins, providing peptides and amino acids as a nitrogen source, which will not be the case for K-DOW, where enzymes have been rendered inactive (Arte et al., 2015; Gänzle et al., 2008). Indeed, prior research showed that there is a major difference in oat endogenous enzymatic activity (amylase and protease) between K-DOW and NK-DOW across the pH range covered during a food fermentation process (Blontrock et al., 2025). Thus, during NK-DOW fermentations, the endogenous amylase and protease activity assisted the inoculated lactic acid bacteria in their growth.

The faster growth of L. plantarum in NK-DOW aligned closely with the higher rate and extent of metabolite production, especially lactic acid (Kedia et al., 2008). During homofermentative lactic acid fermentation, one equivalent of glucose is converted into two lactate equivalents, which become lactic acid in solution. The higher microbial density and higher availability of fermentable saccharides in NK-DOW explain the faster acidification profile of NK-DOW fermentation, reaching pH values below 4.5 after approximately 6 h, compared to K-DOW fermentation, where it took at least 14 h to reach this pH value (Fig. 2A). Thus, from an industrial perspective, the use of K-DOW for fermentation is not recommended, as it may compromise food safety by allowing the potential growth of background pathogens during the initial acidification period (IFT/FDA, 2001). This difference in acidification rate and lactic acid production between K-DOW and NK-DOW corresponded well with the dynamics regarding fermentable saccharide contents. These results are in line with previous studies showing that the addition of extra fermentable saccharides – either directly (Angelov et al., 2006) or through the hydrolysis of oat starch by exogenous amylase (Luana et al., 2014) – accelerates the rate of acidification in kilned oat fermentations.

Besides lactic acid, the increase in the content of other metabolites, such as glycerol and myo-inositol, was also more pronounced in NK-DOW, compared to K-DOW fermentation (Fig. 5). Passos et al. (2003) demonstrated that L. plantarum MOP-3 (isolated from fermenting cucumbers) can convert malic acid into lactic acid. Altought the current study used a different strain of L. plantarum, a similar meaboltic capability may be present, explaining the gradual decrease of malic acid concentrations during both K-DOW and NK-DOW fermentations. The decrease occurred more rapidly in NK-DOW suspensions, likely due to the higher microbial counts during this fermentation. The increase in myo-inositol content over time in the NK-DOW fermentations but not in the K-DOW fermentations likely results from phytate hydrolysis by endogenous oat phytase rather than from microbial activity. The low concentrations of glycerol in NK-DOW suspensions, followed by a slight increase over the course of fermentation, could be related to a lack of yeast growth during the fermentation process, as glycerol production is commonly associated with yeast activity (Wang et al., 2001). The presence of glycerol in NK-DOW suspensions at 0 h might indicate limited production of this compounds by background microbiota, which are inactivated during kilning, explaining the absence of detectable glycerol concentrations during K-DOW fermentation. Another explanation is the enzymatic hydrolysis of the residual lipids (±2%) into glycerol and free fatty acids by oat lipase (Urquhart et al., 1984). This would also explain why glycerol was not detected during the K-DOW fermentations since oat lipase was deactivated by the kilning process. At 24h, the glycerol content was descreased in the NK-DOW suspensions, likely because the fermenting microbes started utilising glycerol as an alternative energy source following the depletion of sugars after 12h (Doi Y., 2019).

Collectively, these results underscore the critical influence of oat kilning on the fermentative potential of oats, with NK-DOW representing a more dynamic and efficient substrate for safe and efficient microbial fermentation.

4.2. Impact of kilning on the bioconversion reactions of oat constituents during fermentation of oat wholemeal flour

Scientific literature on cereal fermentation frequently emphasises its potential to alter the molecular structure of cereal components to eventually obtain desired features (e.g. flavour, texture, health-related aspects) of the final food product (Gobbetti and Gänzle, 2023; Poutanen et al., 2009; Ye et al., 2024). As previously described for wheat and rye fermentations, most of these characteristics – for example dietary fibre solubilisation, protein solubilisation, and phytate degradation – are achieved through endogenous enzymatic activities, promoted by the pH change associated with a typical fermentation process. Such enzymatic conversions have thus far been poorly investigated for oat-based fermentations. Therefore, we systematically investigated the effect of lactic acid fermentation on enzyme-mediated changes in the chemical structure of proteins, dietary fibres, and phytate in oat wholemeal flour and examined how these changes are affected by the prior groat kilning process.

Lactic acid fermentation of DOW had a clear impact on the extractability of dietary fibres (β-glucan and arabinoxylan) (Fig. 6A and B). Fermentation of K-DOW suspensions displayed a consistent increase in β-glucan extractability over time, while arabinoxylan showed a similar but less pronounced trend. Since K-DOW lacks active endogenous oat β-glucanase and endo-xylanase due to kilning, these increases are most likely caused by a better hydration of dietary fibre molecules over time (Blontrock et al., 2025). The overall higher extractability of oat β-glucans compared to arabinoxylan were anticipated since β-glucans are inherently more water-soluble (Flander, 2012). During fermentation of NK-DOW suspensions, fibre solubilisation was much more pronounced, likely resulting from oat endogenous enzyme activity. In line with Blontrock et al. (2025), where it was shown that endo-xylanases are optimally active at pH 4.5-6.0 in non-kilned oats, most arabinoxylan solubilisation likely occurred during the first 6 h of fermentation, as the pH decreases from 6.3 to 4.0 within this period (Fig. 2A). The 2.5-fold increase in AX extractability in NK-DOW suspensions after 24 h was similar to that previously reported for wheat sourdough fermentations (Coda et al., 2014; Korakli et al., 2001; Lappi et al., 2010a, Lappi et al., 2010b). The evolution of β-glucan extractability demonstrated a different profile compared to what was seen for AX, suggesting that insoluble β-glucans were solubilised early in the fermentation process, after which these molecules were further degraded into very low molecular weight compounds by the action of oat endogenous β-glucanases. Earlier research showed that these enzymes have an optimal activity at pH 5.5 but also remain active at lower pH values (Blontrock et al., 2025). The predominant β-glucanases found in oats are (1-3,1-4)-β-glucanases which hydrolyse β-glucans at internal glycosidic linkages, resulting in the production of low molecular weight (1-3,1-4)-β-D-glucan and oligosaccharides (Perrot et al., 2022; Yun et al., 1993). To date, the production of β-glucanases by lactic acid bacteria has not been reported in the literature (Jin et al., 2023). β-glucosidases, which act on terminal non-reducing β-D-glucosyl residues, may further degrade these (1-3,1-4)-β-D-glucan fragments into glucose monomers. Although β-glucosidase activity has been reported in numerous L. plantarum strains, it remains unclear whether the strain used in this study also produced this enzyme during fermentation (Paventi et al., 2025). In any case, the low molecular weight β-glucans, either β-D-glucan oligosaccharides or free glusose, probably did not precipitate in 90% v/v ethanol and were therefore not detected as β-glucans in the used method, as also explained by Blontrock et al. (2025). A similar conclusion was made by Lu et al. (2019) for an oat sourdough (1:1 flour:water ratio) fermented for 12 h with a L. plantarum strain.

The substantial increase in the ratio of free/total phosphate in the NK-DOW fermentations, together with the gradual increase in myo-inositol content in the NK-DOW fermentations, suggests that a portion of oat phytate was completely hydrolysed into myo-inositol and phosphate (Fig. 6C). While this has not yet been reported for oat, complete hydrolysis of phytate by wheat phytase has been reported (Bohn et al., 2007). The minimal decrease of phytate in the K-DOW suspensions after 24 h could be explained by consumption of free phosphate by the fermenting lactic acid bacteria (Gobbetti and Gänzle, 2023). Although multiple studies have reported microbial phytase activity in L. plantarum strains (Haros et al., 2008; Nuobariene et al., 2015; Sandez Penidez et al., 2020; Sumengen et al., 2013), it seemed that no phytase hydrolysis occurred in K-DOW suspensions. This might suggest that the strain used in this study lacks the genetic capacity to produce phytases or that the fermentation conditions did not favour the production of the enzyme. Previous studies showed that incubation of NK-DOW suspensions with gradual acidification and in the complete absence of microorganisms already achieved 41% of free/total phosphate ratio, indicating that endogenous oat phytase is predominant over microbial phytase in hydrolysing phytate (Blontrock et al., 2025). Nevertheless, the obtained free phosphate content in fermented NK-DOW suspensions remained relatively low and – assuming all measured phytate was present exclusively as myo-inositol hexakisphosphate – corresponds to an estimated phytate reduction of 28%. Comparable, low values of phytate hydrolysis of 16-45% during oat fermentation have also been reported by Reale et al. (2007) and Bartnik and Szafrańska (1987). Much higher phytate reductions (70-90%) have been shown for wheat fermentations (Leenhardt et al., 2005; Lopez et al., 2001; Türk et al., 1996). This propbaly relates to inherently low endogenous phytase activity in oat compared to other cereals, as demonstrated by Mayer et al. (2023) and Steiner et al. (2007). Therefore, there is still a clear potential to further increase phytate degradation in oats during fermentation. Further research should focus on selecting starter cultures with high microbial phytase activities or on adding other sources of exogenous phytase.

Fermentation is claimed to alter the molecular structure of proteins, thereby potentially enhancing their extractability. However, for the variety of oat wholemel flour used in this study, this effect was limited as fermentation of NK-DOW suspensions resulted in only a modest increase in protein extractability (Fig. 6D). SE-HPLC analysis showed that this increase is primarily attributable to the formation of small peptides (and free amino acids), while the oat globulin fraction remained largely unaffected. In contrast, no improvement in protein extractability was found for K-DOW fermentations, suggesting that the endogenous proteolytic activity present in NK-DOW is the key driver of the changes therein. In line with this, the only study investigating the fermentation of kilned oat bran reported no significant hydrolysis of oat globulins and even a reduction in protein solubility due to acidification (Loponen et al., 2007).

In this study, oat wholemeal flour derived from non-kilned oat groats was used to enhance oat fermentability. However, such systems are more susceptible to rancidity development because endogenous lipases remain active, indicating a potential trade-off between fermentability and rancidity. To mitigate this risk, the raw materials were defatted before fermentation. Interestingly, previous studies have shown that oat lipase is completely inhibited at pH values below 5.5 (Blontrock et al., 2025; Ekstrand et al., 1992). This suggests that non-kilned oats may be suitable for food products with a low pH, such as fermented products. Future research should therefore investigate the impact of fermentation on rancidity development and sensory properties in non-kilned oat-based foods.

5. Conclusion

To conclude, the results of this study reveal that the prior heat-treatment of oat groats has a strong impact on the fermentability of oat wholemeal and associated biochemical conversion of starch, protein, fibre, and phytate. Fermentation of NK-DOW suspensions significantly improved the solubilisation of dietary fibres and led to partial phytate degradation and modest protein breakdown. Such conversions may well be important from both technological as well as nutritional points of view in many food products such as oat-based sourdough breads or oil-in-water emulsions. Further research should examine how fermentation influences the oxidative stability of non-kilned oat wholemeal flour. Furthermore, enzyme-assisted oat fermentation and fermentation using carefully selected microbial strains hold potential to further enhance protein extractability and increase phytate degradation in oats. Finally, finetuning the oat groat heat-treatment conditions could be explored as a way to selectively inactivate lipase, thereby improving oxidative stability, while retaining sufficient amylase and protease activity to assure a good fermentation quality.

Credit author statement

Eline Lambrechts: Data curation, Methodology, Investigation, Formal analysis, Writing – original draft, Visualisation, Validation.

Ewoud Blontrock: Data curation, Methodology, Investigation, Formal analysis, Writing – original draft, Visualisation.

Thomas Gettemans: Data curation, Methodology, Investigation, Formal analysis, Writing – original draft, Visualisation.

Jolien Lemoine: Investigation, Formal analysis.

Sarah Vanhove: Investigation, Formal analysis.

Yamina De Bondt: Writing – review & editing, Supervision, Methodology, Conceptualization.

Frederik Janssen: Supervision, Methodology, Conceptualization.

Stefan Weckx: Writing – review & editing, Supervision, Resources, Methodology, Funding acquisition, Conceptualization.

Luc De Vuyst: Resources, Methodology, Funding acquisition, Conceptualization.

Arno G.B. Wouters: Writing – review & editing, Supervision, Resources, Methodology, Funding acquisition, Conceptualization.

Christophe M. Courtin: Writing – review & editing, Supervision, Resources, Methodology, Funding acquisition, Conceptualization.

Declaration of generative AI and AI-assisted technologies in the manuscript preparation process

During the preparation of this work the authors used ChatGPT (OpenAI GPT-5, CA, USA) in order to enhance the language quality. After using this tool/service, the authors reviewed and edited the content as needed and take full responsibility for the content of the published article.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

The authors acknowledge financial support from the project HealthFerm, which is co-funded by the European Union under the Horizon Europe grant agreement No. 101060247 and the Swiss State Secretariat for Education, Research and Innovation (SERI) under contract No. 22.00210. Views and opinions expressed are however those of the author(s) only and do not necessarily reflect those of the European Union or European Research Executive Agency (REA). Neither the European Union nor REA can be held responsible for them.

Frederik Janssen and Yamina De Bondt gratefully acknowledge the Research Foundation – Flanders (FWO Vlaanderen, Brussels, Belgium) for a position as postdoctoral researcher (grant numbers 1224123N and 12B3723N, respectively). Eline Lambrechts would also like to thank the Research Foundation – Flanders (FWO Vlaanderen) for providing her with a doctoral grant (grant number 1S43725N). Christophe Courtin and Arno Wouters further acknowledge FWO Vlaanderen for providing funding for the research project with grant number G049824N.

Footnotes

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.crfs.2026.101349.

Appendix A. Supplementary data

The following are the Supplementary data to this article:

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Data availability

The sequenced reads are available at the European Nucleotide Archive of the European Bioinformatics Institute (ENA/EBI) under the BioProject accession number PRJEB102265. Other data is available via https://doi.org/10.5281/zenodo.17550221.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Multimedia component 1
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Data Availability Statement

The sequenced reads are available at the European Nucleotide Archive of the European Bioinformatics Institute (ENA/EBI) under the BioProject accession number PRJEB102265. Other data is available via https://doi.org/10.5281/zenodo.17550221.

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