Borrelia afzelii Infection in the Rodent Host Has Dramatic Effects on the Bacterial Microbiome of Ixodes ricinus Ticks

Abstract

The microbiome of blood-sucking arthropods can shape their competence to acquire and maintain infections with vector-borne pathogens. We used a controlled study to investigate the interactions between Borrelia afzelii, which causes Lyme borreliosis in Europe, and the bacterial microbiome of Ixodes ricinus, its primary tick vector. We applied a surface sterilization treatment to I. ricinus eggs to produce dysbiosed tick larvae that had a low bacterial abundance and a changed bacterial microbiome compared to those of the control larvae. Dysbiosed and control larvae fed on B. afzelii-infected mice and uninfected control mice, and the engorged larvae were left to molt into nymphs. The nymphs were tested for B. afzelii infection, and their bacterial microbiome underwent 16S rRNA amplicon sequencing. Surprisingly, larval dysbiosis had no effect on the vector competence of I. ricinus for B. afzelii, as the nymphal infection prevalence and the nymphal spirochete load were the same between the dysbiosed group and the control group. The strong effect of egg surface sterilization on the tick bacterial microbiome largely disappeared once the larvae molted into nymphs. The most important determinant of the bacterial microbiome of I. ricinus nymphs was the B. afzelii infection status of the mouse on which the nymphs had fed as larvae. Nymphs that had taken their larval blood meal from an infected mouse had a less abundant but more diverse bacterial microbiome than the control nymphs. Our study demonstrates that vector-borne infections in the vertebrate host shape the microbiome of the arthropod vector. IMPORTANCE Many blood-sucking arthropods transmit pathogens that cause infectious disease. For example, Ixodes ricinus ticks transmit the bacterium Borrelia afzelii, which causes Lyme disease in humans. Ticks also have a microbiome, which can influence their ability to acquire and transmit tick-borne pathogens such as B. afzelii. We sterilized I. ricinus eggs with bleach, and the tick larvae that hatched from these eggs had a dramatically reduced and changed bacterial microbiome compared to that of control larvae. These larvae fed on B. afzelii-infected mice, and the resultant nymphs were tested for B. afzelii and for their bacterial microbiome. We found that our manipulation of the bacterial microbiome had no effect on the ability of the tick larvae to acquire and maintain populations of B. afzelii. In contrast, we found that B. afzelii infection had dramatic effects on the bacterial microbiome of I. ricinus nymphs. Our study demonstrates that infections in the vertebrate host can shape the tick microbiome.

Keywords: Borrelia afzelii; Ixodes ricinus; Lyme disease; dysbiosis; egg surface sterilization; microbiome; tick-borne disease; vector-borne disease.

FIG 1

Egg surface sterilization decreases the abundance of the bacterial microbiome in I. ricinus larvae. Each of 10 egg clutches was split into two batches; one batch was surface sterilized (10% bleach for 5 min, 70% ethanol for 3 min) to create dysbiosed larvae, and the other batch was bathed in distilled water to create the control larvae. At 6 weeks after hatching, duplicate pools of dysbiosed and control larvae were frozen; prior to DNA extraction, one pool was washed with 70% ethanol, and the other pool was not washed (10 families × 2 egg surface treatments × 2 larval pool washing treatments = 40 pools). (A) The pools of 6-week-old dysbiosed larvae (labeled “bleach”) had a ratio of bacterial 16S rRNA to tick calreticulin that was ∼27.5-fold lower than that for the 6-week-old control larvae (labeled “water;” P < 0.001); ethanol washing of the larval pools prior to DNA extraction (P > 0.05) had no effect on the ratio of 16S rRNA to calreticulin genes. (B) Egg surface sterilization, but not washing with ethanol prior to DNA extraction, led to significant shifts in the bacterial community of I. ricinus larvae, as measured by 16S rRNA amplicon sequencing and db-RDA (P < 0.001 and P > 0.05, respectively). The larvae hatching from bleached eggs and water-washed eggs are shown in tan and blue, respectively, whereas for the ethanol washing treatment prior to DNA extraction, unwashed and washed are shown with triangles and circles, respectively. (C) The most enriched taxon in response to egg surface sterilization was OTU2, which was annotated as “Candidatus Midichloria mitochondrii” (P_adj < 0.001). In contrast, the most depleted taxon was OTU23, which was annotated as Pseudomonas. The y axis has units of number of read counts per thousand mapped reads. Colored points in boxplots represent individual data points (pooled larvae, colored by the tick family; N = 10).

FIG 2

Dysbiosing I. ricinus larvae does not affect their vector competence for B. afzelii. Dysbiosed larvae and control larvae were fed on B. afzelii-infected mice and uninfected control mice. Engorged larvae were left to molt into nymphs, which were tested for their B. afzelii infection status and bacterial abundance at 4 weeks after the larva-to-nymph molt. For panels A and B, vector competence is only shown for the subset of nymphs that fed as larvae on the B. afzelii-infected mice; the nymphs that fed as larvae on the uninfected control mice are not shown. (A) The percentages of nymphs that acquired B. afzelii during their larval blood meal were similar between the control group (labeled “water”) and the dysbiosed group (labeled “bleach”) (P > 0.05). (B) The nymphal spirochete loads, measured as the ratio of the Borrelia flagellin gene to the tick calreticulin gene, were similar between the control group (labeled “water”) and the dysbiosed group (labeled “bleach”) (P > 0.05). For panels A and B, there was no effect of ethanol washing prior to DNA extraction of the nymphs on vector competence (all P > 0.05). (C) There was a significant interaction between B. afzelii infection status of the mouse and ethanol washing of the dead nymphs prior to DNA extraction. Feeding on B. afzelii-infected mice decreased the bacterial load in I. ricinus nymphs that were washed with ethanol prior to DNA extraction (P < 0.001), but the negative effect of B. afzelii infection on the bacterial load was not significant in the unwashed nymphs.

FIG 3

Tick stage and B. afzelii infection affect the tick bacterial microbiome. Heat map of the numbers of reads assigned (log₁₀[x + 1] per thousand) for the top 20 OTUs across all samples in the experiment. Highest taxonomy reliably assigned by Metaxa2 is shown. Dendrograms are based on hierarchical clustering of Bray-Curtis dissimilarities using Ward’s method. The bacterial microbiome of the I. ricinus larvae was less diverse than the bacterial microbiome of I. ricinus nymphs. Larvae had high numbers of Pseudomonas (OTU1) and Rahnella (OTU31), whereas nymphs had high numbers of Sphingomonadales (OTU13) and Midichloria (OTU2). There was a dramatic effect of B. afzelii infection status in the mice on the bacterial microbiome in I. ricinus nymphs. For example, nymphs that fed as larvae on B. afzelii-infected mice had high numbers of Mesorhizobium (OTU6) and Variovorax (OTU35), whereas nymphs that fed as larvae on uninfected mice had high numbers of Stenotrophomonas (OTU5 and OTU30). In contrast, the effects of the egg surface sterilization treatment on the bacterial microbiome of the larvae mostly disappeared from the nymphal stage.

FIG 4

Tick stage and B. afzelii infection affect the tick bacterial microbiome. (A) Compositions of treatment groups and life stages of I. ricinus ticks, with top 40 OTUs of the bacterial microbiome aggregated (as mean of samples per group) at the genus level. (B) As for panel A but at the order level. The bacterial microbiome of the I. ricinus larvae was dominated by Stenotrophomonas (order Xanthomonadales), Pseudomonas (order Pseudomonadales), and Rahnella (order Enterobacteriales). The relative abundance of the genus Midichloria (order Rickettsiales) was higher in the larvae that hatched from surface-sterilized eggs (bleach +) than in the larvae that hatched from the control eggs (bleach −). There is a dramatic change in the bacterial microbiome as larvae develop into nymphs. The relative abundances of two genera, Pseudomonas (order Pseudomonadales) and Rahnella (order Enterobacteriales), decreased dramatically from larvae to nymphs. In contrast, the relative abundances of many other genera increased dramatically from larvae to nymphs, including Stenotrophomonas (order Xanthomonadales), Midichloria (order Rickettsiales), Mesorhizobium (order Rhizobiales), and Variovorax (order Burkholderiales). For the bacterial microbiome of the nymphs, the relative abundance of Stenotrophomonas (order Xanthomonadales) was much higher in the uninfected control group than in the B. afzelii-infected group. In contrast, the relative abundances of the genera Midichloria (order Rickettsiales), Mesorhizobium (order Rhizobiales), and Variovorax (order Burkholderiales) were much lower in the uninfected control group than in the B. afzelii-infected group. In summary, tick developmental stage and B. afzelii infection status of the mouse have dramatic effects on the bacterial microbiome of immature I. ricinus ticks.

FIG 5

Infection with B. afzelii increases the bacterial microbiome diversity in I. ricinus nymphs. (A) Effects of egg surface sterilization and B. afzelii infection on the Shannon diversity index of the bacterial microbiome of I. ricinus nymphs. Nymphs are classified depending on whether they fed as larvae on an uninfected control mouse (green) or a B. afzelii-infected mouse (pink). Infection with B. afzelii in the mouse increased the Shannon diversity of the nymph bacterial microbiome. Comparison of linear mixed-effects models (LMMs) with Akaike’s information criterion (AIC) found that mouse infection status was a stronger predictor than tick infection status of the Shannon diversity of the nymph bacterial microbiome (ΔAIC = 3.441). Points represent individual tick nymphs colored by family of origin. (B) Principal-coordinate analysis (PCoA) of the 16S rRNA counts in the nymphs, colored by experimental factors; B. afzelii infection status in the mouse best stratifies groups on PCoA axis 1 (PCoA1). Comparison of LMMs with AIC found that mouse infection status was a stronger predictor than tick infection status of PCoA1 (ΔAIC = 48.248).

FIG 6

Infection with B. afzelii changes the bacterial microbiome in I. ricinus nymphs. Here, B. afzelii infection status refers to whether the nymphs took their larval blood meal from an uninfected control mouse (“control” group on the x axis) or an infected mouse (“infected” group on the x axis). Of the 40 most abundant OTUs, the relative abundance of 19 OTUs was significantly different (P_adj < 0.05) between nymphs in the uninfected control group and nymphs in the infected group. These 19 OTUs are color coded according to whether they belong to 4 bacterial classes: Alphaproteobacteria (red), Betaproteobacteria (green), Gammaproteobacteria (cyan), or Planctomycetia (purple). The effect of B. afzelii infection differed among bacterial taxa (Fisher’s exact test P = 0.004); the relative abundance increased significantly for many OTUs in the class Betaproteobacteria (green), whereas it decreased significantly for many OTUs in the class Gammaproteobacteria (cyan).

FIG 7

Experimental design of the study. Engorged adult female I. ricinus ticks (n = 10) were collected from roe deer captured in the Chizé forest, France, and laid their eggs in a phytotron. Each of the 10 egg clutches (for simplicity, only 1 egg clutch is shown) was split into two batches; one batch of eggs was surface sterilized (10% bleach and 70% ethanol) and hatched into dysbiosed larvae, and the other batch of eggs received a control treatment and hatched into control larvae. To determine whether the egg surface sterilization treatment reduced the bacterial microbiome, dysbiosed larvae and control larvae were frozen at 6 weeks after hatching for each of the 10 tick families. DNA was extracted from pools of larvae that were either rinsed with ethanol or not prior to DNA extraction (10 families × 2 egg surface treatments × 2 washing treatments prior to DNA extraction = 40 pools of larvae). These 40 pools of larvae were tested for bacterial abundance using 16S rRNA qPCR and for their bacterial microbiome using Illumina amplicon sequencing of the 16S rRNA gene. The remaining larvae for each of the 20 batches (10 tick families × 2 egg surface treatments) were split into two groups of ∼100 larvae (total of 40 groups) that were fed on either an uninfected control mouse (n = 20) or a B. afzelii-infected mouse (n = 20). Engorged larvae were placed in individual Eppendorf tubes and left to molt into nymphs in a phytotron. Four weeks after the larva-to-nymph molt, 9 to 10 nymphs were randomly selected from each of the 40 mice and frozen at −80°C (n = 370 nymphs). DNA was extracted from individual nymphs that were either rinsed with ethanol or not prior to DNA extraction; 356 nymphs provided enough DNA for further analysis. The DNA extractions from these individual nymphs were tested for B. afzelii infection and for bacterial abundance using qPCR assays targeting the flagellin gene and the 16S rRNA gene, respectively. The tick calreticulin gene copy number (estimated via qPCR) was used to standardize the copy numbers of the flagellin gene and the 16S rRNA gene. The bacterial microbiome of the individual nymphs (n = 308) was studied using Illumina amplicon sequencing of the 16S rRNA gene. This figure was created using BioRender.