Shirkey NJ , Manitt C , Zuniga L , Cohen-Cory S .

EY011912 NEI NIH HHS , R01 EY011912 NEI NIH HHS

             axon development

	Figure 1 Netrin influences the forward advancement and morphology of RGC axon growth cones at their target. (A, B) Representative reconstructions of control (A), and netrin‐treated (B), RGC axon growth cones in stage 40 tadpoles expressing EYFP imaged by time‐lapse confocal microscopy over 24 h. White horizontal lines delineate the forward‐back branching boundary used for quantification of axon growth [also see diagram in Fig. 2(A)]. White arrows indicate examples of intrinsic landmarks (bends in the axon) common to all time points that were used to align images. White arrowheads demarcate differences in the branching patterns of growth cones exposed to control vehicle solution or recombinant netrin (see Fig. 2). Illustrated portions of the axons lie completely within the optic tectum, and rostral to caudal (R‐C) orientation of the brain is indicated. Scale bar, 10 μm. (C, D) A measure of rostro‐caudal growth is provided by the change in length of axons exposed to control or netrin treatment over a 24 h period (C), as well as the proportion of axons that lengthened, shortened or remained the same length throughout the observation period (D). Note that while the majority axons in control tadpoles lengthened, axons exposed to netrin‐1 shortened over time or remained the same length. (E, F) The effects of netrin‐1 treatment on RGC axon growth cones of Stage 40 tadpoles were further determined by quantifying the changes in total branch number (E), and total arbor length (F), at each time point relative to time 0. While changes in total branch number and length were similar for axons in control and netrin‐treated tadpoles during the first 6 h of imaging, a delayed effect of netrin is observed on branch number and length by 24 h. Dotted lines indicate within group comparisons; solid lines represent between group comparisons. * signifies p < 0.05, ***p < 0.0005. Error bars indicate SEM.
	Figure 2 Netrin‐1 differentially influences forward versus back branching of simple RGC axons in Stage 40 tadpoles. (A) Diagrammatic representation of the method used to differentiate forward versus back branching. A horizontal dotted line was used to delineate the forward‐to‐back branch boundary on RGC growth cones imaged over time. Asterisks are used to denote stable branches which persisted throughout the observation period; these stable branches, along with distinctive bends and swells in the axon (black arrows), were used as landmarks for image alignment along the forward‐back branch boundary line. (B, D) The difference in the number of total forward (B), or back (D), branches in RGC axons imaged over time was quantified for EYFP‐expressing axons in control Stage 40 tadpoles and in tadpoles injected with netrin‐1 immediately after the initial imaging (time 0). Note the significant effect of netrin treatment on the number of both forward and back branches on RGC axon growth cones 4 h after treatment. Netrin transiently and significantly increased the proportion of back branches between 0 and 4 h relative to controls (D), but prevented the increase in forward branches observed in controls (B). (C) A forward‐to‐back branch ratio provides an additional measure of axon growth. As RGC axon growth cones branched in the optic tectum, they extended a greater portion of forward than back branches, as indicated by a mean forward‐to‐back branch ratio greater than 1. A forward‐to‐back branch ratio of less than 1 as time proceeds is indicative of decreased forward growth relative to the degree of back branching. Solid lines represent between‐group comparisons. *p < 0.05; **p < 0.005, and ***p < 0.0005. Error bars indicate SEM.
	Figure 3 Netrin influences the dynamic branching behavior of recently targeted RGC axons. (A, B) The dynamic behavior of recently targeted RGC axon growth cones in Stage 40 tadpoles is illustrated by the quantification of total branch addition and branch stabilization for every 2‐h interval during the first 6 h after treatment (average 2 h interval), and between 6 and 24 h. (A) Note that the number of new branches added was similar for newly targeted RGC axons exposed to netrin‐1 as for controls during the first 6 h after treatment but was significantly decreased later in the observation period. (B) Analysis of stable branch number reveals that the number of stable branches was similar for control RGC axons and for those exposed to netrin‐1 at all time intervals. (C, D) The relationship between branch addition and stabilization of forward versus back branches was measured for each individual axon and is illustrated by the stacked bar graphs. (C) In proportion, more new forward branches than back branches were added in control axons during every 2 h observation interval, while a similar number of forward versus back branches were added in axons in netrin‐treated tadpoles. Note that, in proportion, significantly more new back branches were added in axons exposed to netrin‐1 than in controls. (D) No significant difference in the proportion of added and stable forward versus back branches was observed between 6 and 24 h after initial imaging for controls or axons exposed to netrin‐1. In (C, D), stacked bars with a value above 100% represent intervals when more of the newly added branches were stabilized than eliminated. Conversely, stacked bars with a total value of less than 100% represent axons where in proportion more branches were eliminated than added at each corresponding time interval. # signifies 0.05 < p < 0.10; *p < 0.05; **p < 0.005, and *** p < 0.0005, ns = nonsignificant versus control. A dashed line represents comparisons within treatment group. Error bars indicate SEM. (E) Diagrammatic representation of the differential branch addition and stabilization in differentiating RGC axon growth cones in control tadpoles versus tadpoles exposed to netrin 24 h after treatment.
	Figure 4 The age and morphological state of the RGC axon affect its branching response to netrin‐1 in vivo. (A, B) Representative images of two distinct RGC axon growth cones (A), and two branched RGC axons (B), in Stage 40 tadpoles at time 0. RGC axon growth cones and branched axons of Stage 45 tadpoles possessed morphologies similar those of Stage 40 tadpoles (not shown). Scale bar, 10 μm. (C, D) The difference in branching rates of RGC axon growth cones (Simple GCs) and branched arbors (Branched axons) of Stage 40 versus Stage 45 tadpoles in control‐treated and netrin‐treated tadpoles is shown graphically by the increase in total branch number over 24 h. C) The increase in branch number (delta total branch number) over a 24 h period is not significantly different for RGC axons in control tadpoles at the two developmental stages examined (Stage 40 and Stage 45), or for RGC axons with simpler or more complex arbors (simple growth cones or branched axons). Note the differential effect of netrin on simple growth cones of Stage 40 tadpoles, and on branched RGC axons of Stage 45 tadpoles, when compared to age‐ and morphology‐matched controls. (D) The difference in the branching rates of RGC axons exposed to netrin is better illustrated by the line graph of data shown in (C). Data on branched RGC axons of Stage 45 tadpoles is as published in Manitt et al., 2009. Dotted lines indicate within group comparisons; solid lines represent between group comparisons. * signifies p < 0.05; **p < 0.005, and *** p < 0.0005. Error bars indicate SEM. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
	Figure 5 DCC receptor blockade interferes with RGC axon growth cone branching in Stage 40 tadpoles. (A, B) The effects of acute anti‐DCC treatment on simple RGC axons of Stage 40 tadpoles 24 h after treatment on rostrocaudal growth is illustrated by the average change in axon length from its initial position at time 0 (A), and by the relative proportion of RGC axons that shortened, lengthened or remained the same length (B). (C–E) The effects of DCC receptor blockade on the branching of RGC axon growth cones is shown by the quantification of the change in total branch number (C), and the change in forward (D) versus back (E), branches from the initial observation period to the next. Note that anti‐DCC treatment interfered with axon branching when compared to controls (D), an effect that was significant on forward branches (D), but not on back branches (E). (F) Axons exposed to anti‐DCC treatment had a forward‐back branch ratio that differed significantly from that of controls by 24 h. (G–J) The effects of anti‐DCC treatment on branch dynamics of RGC growth cones is illustrated by the total number of branches added (G) and stabilized (H), and by the proportion of added and stable forward versus back branches (I, J) during every 2 h observation interval and between 6 and 24 h. Note that in contrast to control and netrin‐1 treatment, anti‐DCC treatment decreased both the addition and stabilization of branches. However in proportion, dynamic changes in branch addition of forward versus back branches were similar to control. *p < 0.05, **p < 0.005, and *** p < 0.0005. Error bars indicate SEM. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
	Figure 6 In culture, RGC axon growth cones respond to acute netrin treatment with increased forward growth. (A, B) Time‐lapse sequence of sample RGC axon growth cones imaged in culture under control conditions (A) and following acute exposure to netrin‐1 (B). Scale bar, 10 μm. (C) The increase in forward advancement of the growth cone from time 0 to subsequent observation intervals is shown by the relative increase in length for each individual axon (time 0 equals 100%). Note that treatment with netrin significantly increased the forward advancement of RGC growth cones in culture 10 min after treatment, in a dose‐dependent manner.
	Figure 7 Dynamic growth cone changes in RGC axons in response to netrin treatment. (A) Representative images from growth cones in culture fixed 1 h post‐treatment, stained with rhodamine phalloidin (red), serve to illustrate the changes in growth cone morphology in response to netrin treatment. The green circle and the green line around the lamellipodial perimeter in the first sample serve to illustrate the areas selected for measurement of lamellipodial area, and area encompassing the growth cone processes, respectively. White arrows indicate filopodial processes in growth cones. (B) Dynamic changes in RGC axon growth cone morphology following netrin treatment are illustrated by this example (time 0). Black arrows point to processes newly extended by the growing axon (see also Fig 8). Scale bar for (A) and (B), 10 μm. (C) Measurements of the lamellipodial area of the growth cone show that within five minutes of netrin treatment the area of the growth cone decreases in size, a behavior that is different from controls. (D) Measurement of the area encompassing the filopodial processes on the growth cone show that netrin treatment gradually decreased the radial spread of the growth cone processes, an effect that became significantly different from controls 50 min post‐treatment. * p < 0.05; **p < 0.005. Error bars indicate SEM.
	Figure 8 Netrin‐1 induces extension of filopodial processes behind the growth cone in RGC axons in culture. (A, B) Time‐lapse sequence of sample RGC axon growth cones imaged in culture under control conditions (A) and following acute exposure to netrin‐1 (B). Black arrowheads point to filopodial processes at the growth cone in (A), and arrows point to processes behind the growth cone in (A) and (B). These representative time‐lapse images illustrate differences in the extension of filopodial processes in control and netrin‐treated (300 ng/mL) growth cones. Scale bar, 10 μm. (C) Quantitation of changes in number of filopodial processes at the growth cone show that netrin treatment results in growth cones with fewer growth cone processes than controls. (D, E) The number of processes behind the growth cone, along the axon shaft (D; back processes), as well as the number of short, spike‐like processes (less than 1.5 μm; (E) behind the growth cone are increased after netrin treatment. (F) Representative growth cones in fixed cultures 1 h post‐treatment, stained with rhodamine phalloidin (red), illustrate the decrease in the number of filopodial processes on the growth cone in RGC axons in cultures exposed to netrin. White arrows indicate back processes. Scale bar, 10 μm. *p < 0.05, and **p < 0.005. Error bars indicate SEM. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
	Figure 9 Acute exposure to netrin‐1 influences presynaptic protein density and DCC expression in RGC axons in culture. (A, B) Representative control and netrin‐treated RGC growth cones in culture immunostained with antibodies to SNAP‐25 (A, presynaptic protein marker) or antibodies to the netrin receptor DCC (B). Retinal explant cultures were grown in culture for 24 h then treated with netrin‐1(100 ng/mL or 300 ng/mL) or vehicle solution for 1 h before fixation and immunostaining. Cultures were also stained with rhodamine phalloidin (red) before antibody staining to visualize the actin cytoskeleton. (A) SNAP‐25 immunoreactivity is observed throughout the axon shaft and at the growth cone in axons in control and netrin‐treated cultures. Note that SNAP‐25 immunoreactivity is increased by the netrin‐1 treatment. Scale bar, 10 μm. (B) DCC immunoreactivity is observed primarily along the axon, and is absent from growth cones in control and netrin‐treated cultures. Dense DCC immunostaining was observed in axons of control cultures but was reduced in axons from cultures treated with netrin‐1 at the two concentrations used. Scale bar, 10 μm. (C) The density SNAP‐25 immunostaining was quantified by counting the number of immunoreactive pixels per total axon length (μm). A dose‐dependent increase in the density of SNAP‐25 immunostaining is observed in growth cones following netrin‐1 treatment. (D) Quantification of the density of DCC immunostaining along the axon showed that DCC levels decrease following exposure to netrin‐1. Sample sizes are indicated by the numerical values inside bars. * signifies p < 0.05 and ***p < 0.0005. Error bars indicate SEM.

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