Seeds of Tifrunner, a normal nodulating peanut cultivar [16] , were treated with 0.1% HgCl₂ solution for 7 minutes with occasional shaking followed by three times of 5-min rinses with deionized distilled-sterilized water (ddH₂O). The treated seeds were soaked in ddH₂O for 48 hours in growth chamber (Percival CU36:5C8, Perry Iowa) in dark at 25˚C, and then rinsed once and transferred to a 12 × 15-inch germination box with a germination paper (SeedBuro Equipment Company, Des Plaines, IL) absorbed 500 ml ddH₂O. Seeds were allowed to germinate for 5 days in the dark growth chamber at 25˚C until tap roots were about 4 - 6 cm long.

Bradyrhizobia strain LB8 [17] was first grown from glycerol stock on YEM plate supplemented with 25 µg/ml chloramphenicol. A single colony from the plate was then grown in YEM broth in 27˚C shaker at 200 rpm until OD 600 = 0.5 - 1.5. Bacteria were precipitated by centrifugation at 1700× g for 10 minutes and washed two times with sterile water. Final pellet was suspended in water and adjusted to OD 600 = 0.12 as inoculum. For inoculation, individual seedling was placed on a petri dish one at a time and 1 ml inoculum of Bradyrhizobia train LB8 was applied slowly over the surface of the seedling root, the excess inoculum was blotted dry at the root tip on a sterile filter paper. The whole inoculation process was performed under sterile condition in a certified clean bench. The seedling was then transferred to a 16 oz sterile SteriCon™-8 tissue culture vessel (Phyto Technology Laboratories). The vessel was pre-laid, at the bottom, with two stacks of the germination paper absorbed with 50 ml of 25% Hoagland solution without nitrogen to promote rhizobia infection. The vessel was sealed with lid and incubated in the dark growth chamber at 25˚C for 3 - 4 days until shoots emerged. Then the box was partially wrapped with aluminum foil to cover the roots and was transferred to a light growth chamber with light/dark cycles of 16 h/8 h, 25˚C/27˚C setting. The culture vessels were opened everyday during microscopic observation and pictures were taken using Olympus MVX10 microscope and microscope-coupled Olympus MVX-TVIXC digital camera by using Picture Frame V3.0 software. Cultivated peanut control seedling roots without Bradyrhizobia train LB8 inoculation, were also observed for root hair development.

At 5 DAG, roots of individual peanut seedlings of both inoculated and non-inoculated groups were observed for appearance of root hair daily for two weeks. Lamp Black (Natural Pigment LLC) water solution was used to mark the secondary roots emerged on 5 DAG or on the day of inoculation or on 0 day after inoculation (DAI). Pictures of peanut roots were taken daily for two weeks using an Olympus MVX-TV1X Cdigital camera and iPAD.

The seedlings of nodulating cultivar Tifrunner were inoculated with Bradyrhizobia strain LB8 at 5 DAG. The top 2 cm root segments close to the hypocotyl region, where not every base of lateral root had RoH, were observed for root hair and nodule development for two weeks to investigate if the presence of root hair is required for nodulation in these top 2 cm root segments. Pictures were taken daily until 16 DAI (i.e. 21 DAG) using an Olympus MVX-TV1XC digital camera and presence and absence of nodule were recorded.

Scanning electron microscopy was performed on peanut root as described by Walter et al. [18] with modifications. Tifrunner seedling roots at 24 hours after inoculation with Bradyrhizobia strain LB8 and prepared for ethanol cryo-fracture SEM. The roots were harvested and immersed into primary fixative containing EM grade, 4% (v/v) paraformaldehyde, 2.5% (v/v) glutaraldehyde in 0.1 M cacodylate buffer, pH 7.3 (Electron Microscopy Sciences). Sections of the root were razor cut into 10 mm pieces and the region of interest near LR1 and LR2 was lightly pre-scored in the direction of fracture plane. Fixation continued under vacuum infiltration and fixed overnight at 4˚C. The roots were rinsed twice in 0.1 M cacodylate buffer, water washed thrice, dehydrated through a gradient of ethanol (25%, 50%, 75%, 95%, 100%) and placed into a parafilm sleeve containing absolute ethanol. The parafilm sleeve was made by cutting a length twice that of the tissue and wrapped into a tube. One end of the tube was crimped closed and filled with absolute ethanol. While submerged under ethanol, the root section was gently introduced into the tube. The last end of the tube was crimped to seal and immersed into LN₂ until bubbling stopped. The frozen parafilm tube rest on a metal block inside a LN₂ filled Styrofoam box and fractured with a pre-cooled razor blade aligned with the pre-scored lines. The two-halves were collected under LN₂ vapor and immediately placed into pre-cooled absolute ethanol. The fractured root pieces were critical point dried (Bal-Tec CPD-030, Leica Microsystems) and mounted with a carbon adhesive tab and graphite paste onto a 12 mm aluminum SEM stub (Ted Pella, Inc.). The samples were further rendered conductive by sputter coating with Au/Pd and argon_(g) (DeskV, Denton Vacuum) and imaged on a Hitachi SU5000 Schottky Field-Emission Variable Pressure SEM (Hitachi High Technologies, America).

Three regions (0.5 - 1 cm, 1 cm - 2 cm and 2 cm above) of the roots were observed for the occurrence of root hair on their lateral roots for three seedlings (or three replicates). The data were analyzed using analysis of variance (ANOVA) to determine the RoH distribution in different segments of the tap root in Excel. The least significant difference (LSD) test was performed to determine the statistically significant differences of distribution of RoH in the tap root segments. A p value < 0.05 was considered as the threshold for statistical significance between different data points.

Roots of individual peanut seedling of both inoculated and non-inoculated were observed daily for the appearance of RoH and the life span of the appeared RoH were monitored daily till they disappeared or senesced. Our observation showed that RoH started to appear at 7 DAG of both inoculated and un-inoculated peanut roots at the base of the lateral roots when the main roots were approximately 6 cm for both transient-thin root hair and densely populated RoH (Figure 1(b) and Figure 2(a), respectively). For the individual peanut roots with RoH, we tracked the life span of the observed RoH from the day after root hair emergence (DARHE) to when they senesced (disappeared from lateral root base). Results showed that once the RoH appeared, it usually took approximately 1 - 3 DARHE to senesce for transient-thin root hair (Figures 1(a)-(f)). However, RoH could stay on the lateral root up to the time of nodule emergence (i.e. 14 DAG) (Figures 2(a)-(d)).

Among the observed roots of nodulating plants, we noticed that not every single lateral root had RoHat their base especially at the upper part (0.5 cm - 2 cm) of the tap root close to the hypocotyl region (Figure 3). Specifically, no or few RoH were observed at approximately 0.5 cm to the hypocotyl region, while the lower part (>2 cm) towards the root tip of the root system had significantly more RoH (Figures 3(a)-(e)). We further tracked lateral roots without RoH for nodule formation with a focus on the lateral roots emerged between 0.5 cm - 2 cm from the hypocotyl region of the main tap root. We observed that a few nodules did form from lateral roots without RoH ever present (Figures 4(a)-4(o)).

Peanut root at 5 DAG, before the emergence of RoH, was inoculated with Bradyrhizobia strain LB8. At 24 hours after inoculation (HAI), when no RoH (normally appear at 7DAG) was observed yet (Figure 5), a cross section of the inoculated peanut root at 0.5 cm - 2 cm top segment of the tap root towards the hypocotyl with lateral roots devoid of root hair was prepared for SEM observation. The observed cross section had two lateral roots with one being emerged after inoculation but before 24 HAI (namely lateral root 1―LR1) and one being emerged before inoculation (namely lateral root 2―LR2). At the bases of both LR 1 and 2, no RoH was observed under SEM. By looking at the base of LR2, no rhizobia accumulation was observed. However, by looking at the base of LR1, impressive patches of rhizobia accumulation were noticed specifically near the surface of a naturally cracked region of the lateral root (Figure 5(b) magnified in Figure 5(c) and Figure 5(d)). A further zoomin, at the surface of LR1 allowed us to observe the accumulation of rhizobia not only on the surface of the epidermal cells but also underneath the biofilm of the root epidermal cells (Figure 5(d)) indicating the penetration of rhizobia through the epidermis of the root. We further zoomed into cortical cells of the main root near the LR1. Surprisingly, rhizobia were found to not just have invaded into the intercellular space of cortical cells (Figure 5(e)), but also occupied within the root epidermal cells (Figure 5(f)).

We found that RoH started to appear at the base of lateral roots at 7 DAG with or without rhizobia infection, indicating that RoH is not induced by Bradyrhizobial inoculation. However, Wissuwa and Ae [20] observed root hair on 14-day-old peanut seedlings. Likely they only reported when significant root hair was observed since the aim of that particular study was to access root hair on 14-day-old peanut seedlings. Otherwise, the difference in day of root hair emergence could be genotype-or cultivar-specific as different genotypes were used in our study and a previous study [20] . RoH life span varied significantly with some fell off at 2 - 3 DARHE and others staying up till nodule formation, normally at 8 DARHE. The variability of the RoH life span may be due to the nature of the

RoH formed. The hairs disappeared at 3 DARHE were transiently-thin root hair while the RoH with extended time of stay were typically thick and dense. We also noticed that the dense RoHs were typically present on actively elongating root regions of the seedlings (from 2 cm region of the tap root downward), while none or few RoHson older root regions (ranging from 0.5 cm - 1 cm top segment of the tap root) during our two-week observation window. This observation is in agreement with previous findings [20] , that there were fewer RoH on older roots when compared to the young actively growing roots. The uneven distribution of the RoH on the root may be caused by uneven hormone distribution along the root during elongation, which are regulating the RoH emergence.

Tifrunner, a normal nodulating peanut cultivar, produces nodules along the roots. However, we noticed that not every lateral root had nodules or RoH at its base, which provided us the opportunity to investigate the relationship between RoH and nodule emergence sites. We observed that there was nodule development from a lateral root base devoid of RoH. This is our first evidence to support that RoH is not required for Bradyrhizobial invasion of peanut root and subsequently not required for peanut nodulation, which is in line with previous observation [21] that the formation of peanut root nodule at the base of the lateral roots was a result of proliferating cell divisions derived originally from the pericycle and not necessarily the presence of RoH. We further solidified that RoH is not required for peanut nodulation by SEM observation of the seedlings’ lateral roots before RoH emergence. The results further confirmed the invasion of Bradyrhizobia in the region of lateral root emergence cells in the absence of RoH at its base.

Karas et al. [22] reported an alternative infection mode “crack entry” in L. japonicus root hairless 1 mutant, while the wild type of L. japonicus is infected through root hair IT infection path. Therefore, they concluded that ‘crack entry’ mode of infection is used in the absence of root hair in L. japonicus. In another word, the root hair is not required for crack entry infection, which is in agreement with our findings, though many other studies indicated that infection of peanut only occurs in the presence of root hair at the base of the lateral roots [7] [13] [14] [15] .

Based on these observations, we hypothesized that the genes controlling nodulation most likely also have pleiotropic effect on the development or appearance of RoH. Pleiotropic effect of genes is not rare cases and there are quite a few reports on this effect. For example, three mlo powdery mildew resistance genes also have impacts on barley yield [23] [24] . Burstin et al. [25] reported that developmental genes have pleiotropic effects on plant morphology and source capacity which eventually have impact on seed protein content and productivity in pea. It has been reported that lateral root emergence, root hair formation, root hair elongation, and nodule development mechanisms have similar phytohormone regulation and auxin being the major positive regulator of these mechanisms [26] . Other hormones such as abscisic acid (ABA), cytokinin, jasmonic acid and so on play opposite roles in these developmental programs/mechanisms [26] . Liang and Harris [27] proposed that the tendency of legumes to form nodules may be linked to a difference in ABA sensitivity. Since root hair formation and nodule development mechanisms in different legume species, M. truncatula and L. japonicas, share some phytohormones [26] , it is highly possible that peanut nodule development and root hair formation mechanisms share the same phytohormones, thus the closely linked or co-segregation relationship observed between nodulation and RoH formation in our study and other reported studies [13] [14] [15] . Further genetic and molecular studies on peanut root hair formation and nodulation need to be conducted to test the relationship between RoH, nodulation, their genetic and hormonal controls.