Chapter 18 Imaging Axonal Transport of Mitochondria

Abstract

Neuronal mitochondria need to be transported and distributed in axons and dendrites in order to ensure an adequate energy supply and provide sufficient Ca²⁺ buffering in each portion of these highly extended cells. Errors in mitochondrial transport are implicated in neurodegenerative diseases. Here we present useful tools to analyze axonal transport of mitochondria both in vitro in cultured rat neurons and in vivo in Drosophila larval neurons. These methods enable investigators to take advantage of both systems to study the properties of mitochondrial motility under normal or pathological conditions.

Introduction

Neurons are exquisitely organized and compartmentalized. Their cytoskeleton and motor proteins are necessary to distribute essential components to axons, dendrites, and synapses in sufficient supply and to return organelles and signals from the periphery to the cell soma. Mitochondria are one such important organelle that neurons need to allocate properly. Absent adequate mitochondrial movement, the distal portions of neurons, which in a human can be a meter distant from the cell body, may be deprived of sufficient ATP production and may be incapable of adequately buffering cytosolic Ca²⁺. Moreover, the involvement of mitochondria in generating reactive oxygen species and apoptotic signaling heightens the importance of keeping healthy mitochondria in the soma, axons, and dendrites.

Mitochondrial movement is therefore needed to deliver the organelles to the periphery. In addition, it is likely the movement from the periphery back to the soma is needed to provide for protein turnover, since most mitochondrial proteins are nuclear‐encoded and peripheral proteins synthesis is likely to be limited. Damaged mitochondria can be highly destructive to the cell via the production of reactive oxygen species (Hoye et al., 2008). Therefore, turnover of the organelle and refreshment of peripheral mitochondria by fusion with newly minted mitochondria that arrive from the soma may be an important means of maximizing mitochondrial function and minimizing their deleterious potential. This may be one of the most significant reasons that 30–40% of the mitochondria in an axon or dendrite are in motion at any given time (Chen et al., 2007, Overly et al., 1996, Waters and Smith, 2003). Moreover, the optimal density of mitochondria differs from one neuron to another, even from one synapse from another within the same neuron, and is constantly changing (Hollenbeck and Saxton, 2005) as the activity of the neuron shifts the energetic needs of a subcellular domain. Activation of receptors and channels can also place an acute burden of Ca²⁺ influx on a subcellular region, and this too may require a shift in mitochondrial distribution. The necessity of tailoring mitochondrial density to changing needs may be an additional reason that mitochondria are so frequently in motion.

Mitochondria can congruently move a long distance without stopping, or travel a short distance and stop and start again. The majority of their movements are microtubule‐based, either anterograde via kinesins, toward the (+)‐ends of the microtubules such as in axonal terminal or retrograde via dynein, toward the (−)‐ends in the cell bodies (Chang et al., 2006, Hollenbeck and Saxton, 2005, Overly et al., 1996). It is not surprising, therefore, that the dynamic nature of mitochondria is significant for neurons to maintain neuronal function and plasticity, and the abnormalities in mitochondrial distribution have been found in many neurodegenerative disease models such as Charcot–Marie–Tooth (Baloh et al., 2007), Amyotrophic Lateral Sclerosis (De Vos et al., 2007), Alzheimer's (Pigino et al., 2003, Rui et al., 2006, Thies and Mandelkow, 2007), Huntington’s (Trushina et al., 2004), and Hereditary Spastic Paraplegia (Ferreirinha et al., 2004, McDermott et al., 2003). To understand the means by which mitochondrial distribution is achieved and the dynamic regulation of that distribution, static imagery is insufficient. Only through live video microscopy can the mechanisms governing mitochondrial distribution be understood.

Although great progresses have been made in visualizing the transport of neuronal mitochondria, there remain significant challenges that include finding appropriate mitochondrial markers, an ideal perfusion/imaging system to keep neurons alive, and an amenable in vivo animal model. Offsetting these difficulties, neurons also offer advantages over other cell types for studying basic mechanisms of mitochondrial movement. The long parallel arrays of microtubules in neurites provide a simpler geometry in which long‐distance movements of mitochondria can be followed. In addition, axons have largely uniform polarities of microtubules, with all (+)‐ends oriented toward axon terminals. Therefore, the study of mitochondrial transport in axons provides the opportunity to distinguish kinesin‐based movements from dynein based. In dendrites, microtubule polarity is typically more mixed, but the analysis of dendritic motility offers the potential to examine how that motility is affected by the activation of postsynaptic receptors on the dendrite surface and the association of mitochondria with postsynaptic specializations.

Here we describe two systems that have been used successfully in our laboratory and elsewhere to study axonal transport of mitochondria: cultured rat hippocampal neurons and Drosophila neurons. Both systems permit detailed measurements of the parameters of mitochondrial movement and the manipulation of the mechanism either via transgene expression or classical genetics. Separately, they provide the complementary advantages, detailed below, of either in vitro tissue culture or imaging in a semi‐intact in vivo environment.

Embryonic hippocampal neuronal cultures have been widely used to study protein distribution, neuronal function, and neuronal morphogenesis. Their utility derives from their elegant extension of neurites, their expression of key features of in vivo hippocampal neurons, and their suitability for transfection with transgenes (Kaech and Banker, 2006). Neurons dissociated from rat embryonic hippocampi contain few glial cells and can be cultured in vitro for up to a month. After 3–4 days in vitro, axons have differentiated from dendrites and exhibit unique morphological characteristics, such as long, thin, and uniform diameters, a lack of branching, and obvious growth cones. The identification of axons can be enhanced by transfecting the cells with a fluorescent axonal marker such as synaptophysin‐YFP (Chang et al., 2006), or marking dendrites with a marker such as PSD95‐YFP (Chang et al., 2006).

The axons of cultured hippocampal neurons can reach a few mm in length, enabling the observation of long‐range movements. In these neuronal cultures, it is also easy to apply extracellular agonists, antagonists, toxins, or ionophores to regulate intracellular signaling, change intracellular ion balance, trigger neuronal cell death, or mimic a pathological cellular effect. The consequences of these manipulations on mitochondrial movements can be monitored acutely. These advantages have been exploited extensively for the study of mitochondrial motility (Chada and Hollenbeck, 2003, Chada and Hollenbeck, 2004, Chen et al., 2007, Overly et al., 1996, Waters and Smith, 2003).

In Section 2, we will describe the protocols to analyze mitochondrial movement in cultured rat hippocampal neurons. For optimal tracking of individual mitochondria, we have found it desirable to use cultures that were sparsely transfected with a construct encoding a fluorescent protein fused to a mitochondrial targeting sequence (RFP‐mito). Under these circumstances, the processes of a single transfected neuron can be examined and thus the orientation of movement toward or away from the soma can be determined unambiguously. In densely transfected cultures or when membrane permeant dyes are used to image mitochondria, the bundling of neurites makes it impossible to determine the cell body or origin and the direction of movement. We and others have also found that some mitochondrial dyes may decrease movement of the organelles.

As a model system for the genetic analysis of neurons, Drosophila remains unsurpassed. Existing mutant lines are available for dissecting many cell biological processes and ongoing mutant screens continue to identify additional proteins of interest. Despite progress with RNAi in mammalian systems, a null allele remains the best guaranty of complete loss of function. Large collections of RNAi lines are also available in the fly at low cost with which to screen candidate genes (http://stockcenter.vdrc.at/control/main). Additionally, the ease of combining different mutants and transgenes in an individual organism enables studies of double mutants and phenotypic rescue with modified transgenes in an intact organism. Moreover, many mutations whose early lethality would prevent their analysis in a knockout mouse can be studied in the fly through genetic tricks to make mutations homozygous or express RNAi selectively in small subsets of neurons. In consequence, Drosophila has been used to establish many human neurodegenerative disease models, an endeavor justified by the high degree of similarity of their genome to our own and similarity of the resultant cellular phenotypes to human neurodegeneration (Bolino et al., 2004, Clark et al., 2006, Gunawardena et al., 2003, Wang et al., 2007). Overexpression of a normal or a disease‐related allele can be accomplished by crossing fly strains with cell type or tissue‐specific GAL4 drivers to the fly strains harboring the gene‐of‐interest downstream of a GAL4 binding site (UAS) (Brand and Perrimon, 1993).

For live‐imaging studies, the opaque cuticle of the adult fly poses an optical challenge and makes dissections more difficult. Therefore, most imaging studies have chosen the third‐instar larval stage for analysis. These larvae are easy to dissect and many mutants and transgenic lines of interest survive to this stage. Mutations that severely impair the transport of mitochondria may die before this stage, however; milton null alleles, for example, die in the first instar (Stowers et al., 2002). This need not preclude live imaging of mitochondria, however, though it does make the dissection more challenging and imaging in intact specimens through the cuticle may be preferred by those who have not mastered the dissection.

In Drosophila larvae, the cell bodies of central nervous system neurons form the cortex of the brain lobes and ventral nerve cord (VNC) and are bilaterally symmetrical. In the core of the VNC two parallel zones of neuropil lie to either side of the midline. It is in this neuropil where the axonal and dendritic projections of the cell bodies comingle and communicate. The complexity of the neuropil has, to date, made it unattractive for imaging studies of axonal transport. Instead, laboratories have focused on the nerves that issue from the VNC and extend to the body wall in each segment. These nerves include the axons of motorneurons that emanate from the VNC and terminate in well‐characterized neuromuscular junctions on larval body walls. These segmental nerves also contain sensory axons whose cell bodies reside in the body wall and whose axons extend into the VNC, with the opposite orientation of the motorneurons. Therefore, to know the polarity of mitochondrial movement in these nerves, it is necessary to selectively label a subset of neurons of either the sensory or motorneuron class. To date, this has principally been done by the selective expression in motorneurons of a fusion of GFP with a mitochondrial import signal (Miller et al., 2005, Pilling et al., 2006). In Section 3, we will introduce the protocols to analyze the axonal transport of mitochondria in Drosophila neuronal axons at the late larval stage (third instar) and will describe a variation on the method of Pilling et al. (2006) in which we selectively label mitochondria in one peptidergic axon per segmental nerve.