Trapping, Tagging and Tracking: Tools for the Study of Proteins During Early Development of the Sea Urchin

Abstract

The exquisite synchronicity of sea urchin development provides a reliable model for studying maternal proteins in the haploid egg as well as those involved in egg activation, fertilization and early development. Sea urchin eggs are released by the millions, enabling the quantitative evaluation of maternally stored and newly synthesized proteins over a range of time (seconds to hours post fertilization). During this window of development exist many hallmark and unique biochemical interactions that can be investigated for the purpose of characterizing profiles of kinases and other signaling proteins, manipulated using pharmacology to test sufficiency and necessity, for identification of post translational modifications, and for capturing protein-protein interactions. Coupled with the fact that sea urchin eggs and embryos are transparent, this synchronicity also results in large populations of cells that can be evaluated for newly synthesized protein localization and identification through use of the Click-iT technology. We provide basic protocols for these approaches and direct readers to the appropriate literature for variations and examples.

I. Introduction

A mechanistic understanding of cellular function ultimately depends on studying the native biological materials. New live cell imaging techniques using genetically encoded sensors have revolutionized the field of cell signaling (Oldach & Zhang, 2014; Agetsuma, Matsuda & Nagai, 2017; Ma, Wen, He, Huang, Wang & Zhou, 2017; de la Cova, Townley, Regot & Greenwald, 2017; Lodygin & Flügel, 2017; Ni, Mehta & Zhang, 2018). However, the assessment of protein synthesis, complex interactions, and activity status demands a biochemically tractable system. Studies focusing on cellular signaling require analysis of proteins in the cell, and except for nucleic acid polymers, current technologies are usually incapable of detecting macromolecules from single cells. In such cases, identifying a model system with a high degree of synchrony on a population basis that is also amiable to imaging is highly advantageous.

One of the biggest challenges to studying embryonic development and cell physiology in general is variability from cell to cell, within and between genetically distinct embryo cultures, especially over short time courses. An advantage of the sea urchin (and other echinoderms) is population synchrony - the ability to isolate millions of gametes from a single female which undergo fertilization in a highly synchronous manner proceeding well into early larval development. For example, Tim Hunt and colleagues detected the cell cycle regulatory proteins, the cyclins, in early sea urchin embryos in part because the sea urchin cultures were synchronous in their cell cycles. This started in motion an intense study of cell cycle regulation in the sea urchin leading to a Nobel Prize for discovery of cyclins (Evans, Rosenthal, Youngbloom, Distel & Hunt, 1983; Hunt, 2001). The unparalleled synchronicity of the sea urchin also enables biochemical analysis of egg activation and early embryogenesis on very precise time scales. Resolution on the order of seconds to minutes can be achieved, making the sea urchin egg and embryos especially attractive as a model to study rapid signaling events such as calcium release on a population-based, synchronous basis (c.f. Galione, 2017; Santella, Limatola & Chun, 2015; Morgan & Galione, 2014; Ramos & Wessel, 2013; Tosca, Glass, Bronchain, Philippe & Ciapa, 2012; Townley, Schuyler, Parker-Gür & Foltz, 2009; Roux, Townley, Raisch, Reade, Bradham, Humphreys, 2006; Giusti, O’Neill, Yamasu, Foltz & Jaffe, 2003; Kumano, Carroll, Denu & Foltz, 2001).

In addition to analysis of synchronous populations of cells, the large, optically clear sea urchin eggs and embryos enable excellent imaging capabilities involving manipulations based on microinjection and micromanipulation procedures. These techniques allow for analysis of signaling pathways when properly coupled to in vivo imaging platforms. As more sensitive methods are developed, single cell biochemical approaches are also tenable in echinoderms (Carroll & Hua, 2009; Juliano, Swartz & Wessel, 2014). Select antibodies are sufficiently sensitive to detect proteins in single cells via immunoblotting (Hughes, Spelke, Xu, Kang, Schaffer & Herr, 2014). For example, a single sea urchin egg yields roughly 50 ng of total protein; Carroll and Hua (2009) reported quantitative microinjection of sea star oocytes followed by immunoblotting of single cells using the phospho-specific MAPK antibody.

A challenging aspect of assessing the role of signaling pathways during egg activation at fertilization is the species-specific variation in cell cycle stage with respect to meiotic arrest during oocyte maturation. In most organisms, the arrested oocyte resumes meiosis in response to a maturation hormone, resulting in a fertilization competent egg that then completes meiosis after fertilization (Costache, McDougall, & Dumollard, 2014; Sagata, 1996; Nebreda & Ferby, 2000). In contrast, the sea urchin spawns eggs that have completed meiosis and are naturally fertilized in a haploid stage. They are poised to directly enter the first mitotic cell cycle at fertilization; therefore, they provide an excellent system in which to study changes in protein dynamics in response to the fertilization event independently of meiotic maturation events.

This chapter provides an overview of the main considerations for experimental design and basic methods for investigating rapid, endogenous signaling events and changes in native protein status in a synchronous system. These include assessing the activity of cell signaling pathway components (via targeted and proteomic approaches), nascent protein synthesis using Click-iT technology, and activity through immunoblotting and (phospho) proteomics in the sea urchin. These methods and consideration can be adapted for other echinoderms and beyond.

II. Using the Synchronous Sea Urchin System to Investigate Cell Signaling

Many resources exist for the collection and culturing of sea urchin eggs and early embryos (Foltz, Adams & Runft, 2004; see also the companion Volume 150, Part A, Chapter 1). However, these protocols require extra care to ensure synchronicity when accurate and quantitative biochemical measurements of cell signaling pathways is desired. In general, minimize the processing time between collection and lysis of samples, rigorously document the uniformity and health of cultures, and pay careful attention to accurate timing, temperature and handling of eggs, embryos and lysates. As a result, it is practical to consider employing an assembly line method for sample processing and to carry out trial runs to optimize synchronous fertilization and sample processing logistics.

A. Methods and considerations for obtaining and evaluating synchronicity of a culture

1. Gametes are collected from adult sea urchins induced to spawn by KCl injection (Foltz et al, 2004; Volume 150, Part A, Chapter 1). Handle with care to reduce potential disturbances to cellular signaling and artificial induction of egg activation. Avoid prolonged periods of settling and when resuspending, use a paddle with gentle stirring. During gamete spawning and collection, maintain the eggs within the species-specific temperature range, ~12^oC for S. purpuratus. This can be accomplished using pre-chilled reagents and glassware in a temperature controlled facility or water bath with constant monitoring. Temperature maintenance is important as minor changes in cell signaling, occurrences of artificial induction of egg activation or parthenogenesis, and elevated temperature all have the potential to reduce synchronicity of cultures. Avoid combining eggs from different females, as this also reduces synchronicity.

2. Release of eggs should occur directly into pre-chilled filtered (0.22 μm) natural sea water (FSW). Washes and de-jellying protocols should be carefully considered based on experimental aims. A gravity wash performed 3 times using pre chilled FSW (filtered, 0.22 μm) followed by two gravity washes in artificial sea water (ASW- 484 mM NaCl, 10 mM KCl, 27 mM MgCl₂, 29 mM MgSO₄, 11 mM CaCl₂, 2.4 mM NaHCO₃, pH 8.0; see Volume 150, Part A, Chapter 1 for seawater considerations) is required for removal of contaminating microorganisms as well as decanting of unhealthy eggs and immature oocytes. Successful de-jellying by passage 8 times through 120 μm Nitex mesh (Barnstead, Inc; Barnstead, NY) is encouraged to remove the extracellular jelly coat. Removal of jelly (i) reduces extracellular protein contamination in subsequent biochemical assays; (ii) increases the ability to control the sperm acrosome reaction using jelly water:ASW (1:1 mix) and; (iii) decreases egg buoyancy leading to faster settling times during washes, reducing processing time. Although washed, normoxic sea urchin eggs can be used over a span of hours, minimizing the time (~2 hours maximum) between spawning and fertilizing greatly enhances synchronicity.

3. Assess the spawned eggs by microscopy (10X objective) immediately and during every subsequent gravity wash. The following conditions must be met for an individual batch of eggs to be considered for a synchronous culture (i) eggs lack a germinal vesicle; (ii) cytoplasm is uniformly pigmented (light orange to dark brown in S. purpuratus); and (iii) diameter of egg measures within 5% of published range for the species (see Volume 150, Part A, Chapter 1; ~80 μm for S. purpuratus). Each aspect should exist uniformly throughout a single spawned culture, from a single urchin, with less than 5% variation. To avoid inclusion of spontaneous or parthenogenic activation, avoid using batches of eggs that exhibit any fertilization envelope (FE) elevation prior to fertilization.

4. Sperm should be collected dry by removing it from the animal with a pipette upon spawning and kept on ice. Activation of sperm induced by addition of ASW:jelly water (1:1), should be assessed by light microscopy and determined by presence by vigorous flagellar movement lasting >10 min. A 10% suspension of washed, de-jellied eggs inseminated with a 1:10,000 dilution of activated sperm should result in a 95–100% fertilization success rate within 2 minutes post sperm addition. Fertilization is quantitated by visual confirmation of elevated FE. Monitor FE elevation closely. If < 95% FE elevation is observed, do not use the culture. Depending on the time point desired for biochemical analysis, it is sometimes necessary to begin collecting samples before FE elevation is complete. In this case, one person should be monitoring the culture while another processes the samples. In some cases, removal of the FE may be desired and this adds another step to the procedure (Vol 150, Part A, Chapter1).

5. Once successful FE elevation is observed, and depending on the time point collection schedule, dilute the fertilized culture in a large volume of FSW (10–20 X volume) and wash at least twice by gravity settling to remove excess sperm. Careful consideration should be placed on setting suitable culture density (<1% vol:vol in S. purpuratus), constant temperature and oxygen saturation of embryonic cultures (longer term cultures may require, addition of Penicillin or Streptomycin to prevent bacterial growth). Keeping a log of these and other parameters can help establish the ideal conditions for a given species within the specific laboratory environment. Cultures remain synchronous until planktonic stages are reached although intra-embryo variation of cultures increases during blastogenesis and with the environmental variations mentioned above.

B. Methods and considerations for preparation of non-ionic detergent soluble protein lysates

Total soluble protein from unfertilized eggs, zygotes or embryos can be isolated using a variety of buffers, optimized for a specific experimental question. Using a non-ionic detergent is compatible with many downstream steps and often preserves enzymatic (such as kinase) activity and the ability to carry out (co-) immunoprecipitations. Detailed protocols for immunoprecipitation and protein affinity interactions are provided elsewhere (Roux-Osovitz & Foltz, 2014). All reagents and materials should be prepared fresh and pre-chilled on ice. Protease and phosphatase inhibitors (see below) should be added to buffers immediately prior to use. Here we provide a brief protocol to highlight the key considerations when preparing biochemically stable samples from synchronous cultures. Depending on the time points desired, several people may be needed to collect and process the samples in a timely fashion. Importantly, determine in advance how much total protein is needed for each time point as this will dictate the volume of eggs or embryos collected. In general, buffers containing non-ionic detergents will yield ~ 50–200 mg total soluble protein per 1 mL of a 10% vol:vol culture S. purpuratus eggs or embryos (Roux-Osovitz & Foltz, 2014).

1. Load a pre-chilled tuberculin syringes (1 for each sample) fitted with 27.5 gauge needles with 0.2 mL of the ice cold lysis buffer and place on ice. An example of a multi-use lysis buffer is HNET: 50 mM HEPES pH 7.0, 150 mM NaCl, 15 mM EGTA pH 8.0, 1% NP-40. Add phosphatase inhibitors (0.5 mM Na₃V0₄ sodium vanadate, 1 mM NaF sodium fluoride, 60 mM Na β-glycerophosphate), and a suite of protease inhibitors (10 mM CalBiochem Protease Inhibitor Cocktail III, 50 μM PMSF or PEFABLOC) just prior to use, keeping all reagents on ice.

2. Transfer 1.5 mL of a 10% suspension of eggs or 1% suspension of zygotes/embryos to a pre-chilled microfuge tube and quickly spin in a microfuge (4000 × g) to collect the sample. For faster sample collection, use a mini centrifuge with pulse spin capacity (see Roux-Osovitz & Foltz, 2014). Remove the FSW by pipetting and add the ice cold lysis buffer using the pre-loaded syringe. Re-suspend the eggs and passage the lysate through the needle at least 3 times to lyse (if FEs have been retained and are hardened, more passages may be necessary). Keep the sample on ice at all times and avoid introducing bubbles. A small drop from the needle can be visually inspected using the microscope to assess the extent of lysis. If conducting a time course post fertilization, the time between sperm addition and complete lysis should be noted. This amount of eggs and lysis buffer typically gives a 1:2 pellet:buffer volume ratio. Additional buffer can be added if necessary to aid in lysis. This procedure can be scaled up if large amounts of soluble lysate are required but should be kept on the small scale when multiple tight time points are desired for ease of handling.

3. Incubate the lysates on ice for 10–15 min, followed by centrifugation at 18,000 × g at 4°C for 20 minutes. Carefully transfer the clarified supernatant (the “non-ionic detergent soluble protein lysate”) to a new, pre-chilled 1.5 mL microfuge tube, kept on ice. Insoluble material can be saved and analyzed by direct resuspension in SDS or sample buffer if desired. Soluble lysates should be used immediately or aliquoted and snap frozen in liquid nitrogen then stored at −80^o C. Save a small aliquot for determination of protein concentration. In general, use the lysates immediately until experimental parameters are well established and the effects of freezing are established. Avoid freeze-thawing of the samples.

C. Use of non-ionic lysates in analysis of activity by phospho-specific immunoblotting

C1. General considerations

The egg contains many maternal proteins and mRNAs that are used during oogenesis, immediately following fertilization, or are being stored for use in later in development (cf Picard, Mulner-Lorrilon, Bourdon, Morales, Cormier, Siegl & Bellé, 2016; Chassé, Aubert, Boulben, Le Courguille, Corre & Cormeir, 2018). Thus, it is imperative to identify proteins that are already present in the unfertilized eggs and the post-translational protein modifications that must be occurring to regulate their interactions and activities. For example, the signaling pathways that regulate calcium release at fertilization and the immediate downstream events that propel the egg into the cell cycle rely on post translational modifications (especially phosphorylation) of maternal proteins (Ramos & Wessel, 2013). One example is the MAP Kinase (MAPK) pathway, which is involved in oocyte maturation and arrest as well as cell cycle progression in echinoderms (Kishimoto, 2004). This particular protocol was developed for the detection of the sea urchin MAPK pathway signaling proteins from synchronous batches of eggs, zygotes and embryos providing enough material from each sample for replication of downstream analyses. The MAPK pathway components are highly conserved and antibodies that recognize the proteins in their phosphorylated, active forms, are commercially available. The pathway is therefore an excellent model for investigation of signaling in the sea urchin. Further, the protocols can be adapted to a variety of signaling pathways.

C2. Analysis of the MAPK signaling pathway as a model for phospho-specific immunoblotting

1. Optimal resolution of the MAPK Kinase (MEK) and MAPK proteins via SDS PAGE can be achieved using samples of lysates (20 – 50 μg total protein per lane are easily detected, but sensitivity is adequate for single cells) dissolved in freshly prepared Laemmli SDS sample buffer and heated at 95°C for 10 minutes. Separate proteins by electrophoresis on 10%, 12% or 4–20% gradient polyacrylamide Tris-glycine SDS gels followed by transfer to nitrocellulose membrane. We find that running the gel at a slow voltage (~60V) optimizes resolution. After blocking (5% milk TBST), the membranes are probed and labeled with desired antibodies followed by image capture and analysis (see below).

2. The MAPK pathway antibodies that work well in sea urchins include monoclonal ERK antibody (cat # 610124; BD Transduction Laboratories, Franklin Lakes, NJ), used at 0.25 μg/mL; polyclonal anti-phospho-p44/42 MAPK (Erk1/2, cat # 9101; Cell Signaling; Danvers, MA) used at 1:1000 dilution; and polyclonal anti-MEK 1/2 and polyclonal anti-phospho-MEK 1/2 (cat # 9120 and #9121; Cell Signaling Technologies) used at 1:500. Incubation times of 2.5 hours at RT work well but can be adjusted depending on experimental design. Controls include probing with secondary antibody alone and for normalization, a monoclonal α tubulin antibody (Sigma) used at 1:10000. For direct comparisons, the same blot should be probed with multiple antibodies. In our experience, probing first with anti-phospho-p44/42 MAPK followed by a stripping and reprobing with anti-ERK works best. A quick method for stripping is to rinse the blot quickly in nanopure water followed by a 5 minute incubation in freshly prepared 0.2 M NaOH with moderate agitation. After 4 quick rinses in nanopure water, wash the blot 3 × 5 minutes in block buffer followed by a blocking step and then probe with the next antibody.

3. Multiple methods for immunoblot detection are available. We use secondary antibodies (HRP-conjugated sheep anti-rabbit antibody and goat anti-mouse) from BD Transduction Laboratories used at 1:10,000. Antibody binding is detected using Enhanced Chemiluminescence and imaging using a CD camera system (an example is the BioRad Chemi-Doc platform).

4. Densitometry of bands is calculated using imager-specific software or using open source NIH ImageJ (Wayne Rasband, Research Services Branch, National Institutes of Health, Bethesda, MD; available for Internet download at https://imagej.nih.gov/ij/). Individual sample intensities are first normalized to the internal loading control (tubulin). A measure of activity can be determined by calculating the percentage of phosphorylated MAPK or phosphorylated MEK relative to the total MAPK or MEK (respectively). This can further be assessed for changes over treatments or over developmental time by normalization to the control or unfertilized (reference) time point respectively for each experiment. A minimum of three different experiments must be carefully quantitated, ideally with internal triplicates in each.

D. Methods and considerations for use of pharmacological agents when assessing cell signaling

Egg activation events during fertilization involve potentially redundant and integrated signaling pathways. Formulating a targeted approach to studying specific pathways and components can be informed by employing gain or loss of function conditions with the aid of pharmacological agents. Although off target and toxic effects are always a concern, pharmacological agents can provide the baseline for what pathways to investigate for a given cellular event. This is particularly useful for studying the stable maternal proteins involved in early egg activation, as functional tests that rely on knockout or knockdown strategies may not be feasible.

One of many examples of early signaling is the initiation of sperm-induced egg activation. This can be assessed by evaluating specific physiological events such as internal calcium release and the rise in pH. Calcium ionophores (cf Kumano et al, 2001; Carroll, Albay, Hoang, O’Neill, Kumano & Foltz, 2000) or chelators such as BAPTA (cf López-Godínez, Garambullo, Martinez-Cadena & Garcia-Soto, 2003) work well in echinoderms (see also Bootman, Allman, Reitdorf & Bultnyck, 2018). In addition, many cell signaling pathways can be directly targeted by commercially available pharmacological inhibitors. A good example of this is the MAPK pathway, which can be interrogated using commercially available MEK inhibitors U0126 and PD98059 (Kumano et al, 2001; Carroll et al, 2000; Mulner-Lorillon, Chassé, Morales, Bellé & Cormier, 2017; see Table 1) and the MAPK inhibitor FR180204 (Wang, Sun & Foltz, unpublished). Although a detailed protocol of these methods is beyond the scope of this chapter, the reader is directed to the resources listed in Table 1.

Table 1.

Pharmacological agents (non-inclusive) used effectively in sea urchin eggs and embryos to probe specific signaling pathways.

agent	function/target1	effective final concentration	commercial source	reference(s)
U73122	phospholipase C	5–25 μM	Calbiochem #662035	Lee & Shen, 1998
pertussis toxin (A subunit)	Gαi	0.2 μg/mL	Sigma #P7208	Voronina & Wessel, 2004
Mastoparan-7	Gαi (activator)	10 μM	Biomol #LKT MO172	López-Godínez et al, 2003; Voronina & Wessel, 2004
PD98059	MEK	2.5 μM	Promega #V1191	Carroll et al, 2000
U0126	MEK	2 – 20 μM	Promega #V1121	Houel-Renault et al, 2013; Kumano et al, 2001; Mulner-Lorillon et al, 2017; Kisielewska et al, 2009
FR180204	MAPK (ERK)	0.25 μM	Tocris #3706	Wang, Sun & Foltz, unpublished
BRD 7389	RSK	5 μM	Tocris #4073	Kurylo & Foltz, unpublished
PP1	Src type kinases	10 μM	Tocris #1397	Abassi et al, 2000
API-2	AKT/protein kinase B	2–25 μM	Tocris #2151	Robertson et al, 2013
SB216763	GSK3	1 μM	Tocris #1616	Robertson et al, 2013
PF 573228	Focal Adhesion Kinase	10 μM	Tocris #3239	Chan et al, 2013
Y11	Focal Adhesion Kinase	2.5 mM	Tocris #4498	Chan et al, 2013
roscovitine	CDKs	20 μM	Sigma #R7772	Houel-Renault et al, 2013; Kisielewska et al, 2009; Chan et al, 2013
PP242	mTOR kinase	10–30 μM	Calbiochem #P0037	Chassé et al, 2016
wortmannin	PI3 kinase	20 μM	Sigma #W1628	Houel-Renault et al, 2013; Bradham et al, 20042
LY294002	PI3 kinase	20 μM	Cell Signaling #9901	Houel-Renault et al, 2013; Bradham et al, 20042
bafilomycin A1	V-ATPase	100 nM	Sigma #B1793	Houel-Renault et al, 2013

¹Inhibits the function of the target unless otherwise indicated.

²These authors indicate that the inhibitors were effective at lower concentrations and that the inhibitors slowly inactivated over time diluted in DMSO and stored frozen.

Key considerations include: (i) determining solubility of the pharmacological agent in ASW; (ii) whether it can be taken up by fertilized eggs that retain the FE; (iii) the developmental window of exposure; (iv) length of time of exposure; (v) whether the agent can be washed out; and (vi) dose-dependency. Negative controls such as mock treatment and treatment with vehicle alone are critical. In some cases, inactive analogs of pharmacological reagents are available and serve as excellent negative controls. An example is the MEK inhibitor U0126 and the inactive analog U0124 (Kumano et al, 2001; Carroll et al, 2000; Mulner-Lorillon et al, 2017). Whenever possible, conduct experiments in triplicate and assess whether results are replicated in genetically distinct batches of eggs (from at least three different females). Readers are directed to Vol 150, Part A, Chapters 17 and 18 of further considerations of best practices for studies of abiotic stressors on echinoderm embryos and larvae. Finally, whenever possible, pharmacological results should be validated through the use of specific, targeted approaches such as antisense knockdown (cf Robertson, Coluccio, Jensen, Rydlizky & Coffman, 2013; Oulhen, Swartz, Laird, Mascaro & Wessel, 2017), antibody inhibition (cf Voronina & Wessel, 2004; Schumpert, Garcia, Wessel, Wordeman & Hille, 2013) or dominant inhibition (cf Abassi, Carroll, Giusti, Belton & Foltz, 2000; Chan, Thomas & Burke, 2013).

III. Isolation of proteins for global proteomics, phosphoproteomics and identification of protein complex association

A. Advance considerations

Isolation of healthy gametes and generation of synchronous cultures (see Section II) are necessary starting points for the successful preparation of proteins for use in proteomic studies. An advantage of the sea urchin system is that a single female yields enough gametes to set multiple cultures for large scale sampling and internal replication. In recent efforts (Guo, Garcia-Vedrenne, Isserlin, Lugowski, Morada & Sun, 2015; Wan, Borgeson, Phanse, Tu, Drew & Clark 2015), identification of sea urchin egg and early embryo proteins via high throughput, quantitative proteomics was achieved using triplicates of four different crosses (i.e., 12 samples total representing triplicates of 4 different females), allowing for robust statistical analysis. Earlier work (Brandhorst, 2004; Roux et al, 2006, 2008; Roux-Osovitz & Foltz, 2014) using 2-dimensional electrophoresis provided the essential foundation and identified key parameters for the high throughput methods. The method outlined here was optimized to preserve protein-protein interactions as well post translational modifications (PTMs) such as phosphorylation. In these experiments, eggs are gently homogenized in an intracellular-like buffer and the pH is adjusted to mimic the unfertilized egg (pH 6.9) or fertilized egg (pH 7.4). The buffer composition was based on published methods developed to assess enzymatic activity in cell free lysates (Winkler & Steinhardt, 1981; Zhang & Ruderman, 1993), but can be adjusted to address desired activities or PTMs. Morgan and Galione (2014) describe a similar homogenization buffer optimized for assaying Ca²⁺ and other second messenger mobilization. As with any protein preparation, careful attention to minimizing proteolysis and maintaining PTMs such as phosphorylation are critical.

In addition to preserving PTM of proteins, close attention should be paid to downstream protein analysis workflow compatibility. Depending on the goal, methods used for solubilization are especially important, since detergents may interfere with the proteomic analysis workflow (Vuckovic, Dagley, Purcell & Emili, 2013). Solubility is the main consideration if the purpose of the experiment is the isolation of individual proteins for analysis of changes in quantity or modification over time (Guo et al, 2015). However, if the goal is to capture unbiased protein-protein interactions via a co-fractionation approach (Wan et al, 2015), careful attention to both detergent and ionic strength parameters are important in order to optimize solubility while maintaining interactions. Ultimately, the proteins will be digested and resulting peptides subjected to Time of Flight Tandem Mass Spectrometry (ToF MS/MS). Therefore, pilot experiments are necessary to determine optimal methods for solubilizing proteins and digestion of the samples. Finally, since methods are well-established for isolating organelles and other components of sea urchin eggs and embryos (cf Wessel & Vacquier, 2004), recent advances in analyzing organellar and membrane proteins in high throughput proteomic platforms (cf Vuckovic et al, 2013; Kar, Simonian & Whitlegge, 2017) should be adaptable to the sea urchin system as well.

B. Detailed protocol for the isolation of proteins for global proteomics, phosphoproteomics and identification of protein complex association

B1. Materials Needed

These methods have been successful for both S. purpuratus and L. pictus. All buffers, vessels, sample tubes, centrifuges and rotors should be pre-chilled to 4^o C. If conducting a tight time course, practice how long it takes to process a given sample and enlist help as needed to accomplish the temporal constraints. In the method described here, we routinely capture 2- and 5 minutes post sperm addition time points (time between adding sperm and complete lysis). Buffers should be freshly prepared using ultrapure water and pre-chilled. Typically, 500 mL of each buffer is enough for a large scale preparation. Add protease and phosphatase inhibitors just before use to small quantities of buffer to ensure stability. The ASW recipe is provided in Section II A.2 (above).

1. Buffer G (pH 6.9): 40 mM NaCl, 2.5 mM MgCl₂, 300 mM glycine, 100 mM potassium gluconate, 2% glycerol, 50 mM HEPES, 10 mM EGTA, 4.19 mM CaCl₂. The pH of the chilled buffer should be adjusted using KOH.

2. Buffer G (pH 7.4): 40 mM NaCl, 2.5 mM MgCl₂, 300 mM glycine, 100 mM potassium gluconate, 2% glycerol, 50 mM HEPES, 10 mM EGTA, 8.56 mM CaCl₂. The pH of the pre-chilled buffer should be adjusted using KOH.

3. Just prior to use, add protease and phosphatase inhibitors to the required volume of buffers to achieve the following final concentrations:

   • 1 μM PEFABLOC (a PMSF mimic, Roche)
   • 10 mM Protease Inhibitor Cocktail 3 (Cal Biochem) or Complete inhibitor cocktail (Roche).
   • 1 mM Na orthovanadate and 100 mM NaF or PhosStop (Roche)

4. A protein concentration determination assay of choice (such as the Pierce Endogen BCA Assay).

B2. Isolating soluble proteins from eggs and embryos

1. Collect gametes by intracoelomic injection of 0.55 M KCl and de-jelly the eggs. After washing, suspend the eggs at 10% vol:vol in ASW. Remove an unfertilized egg sample and set aside for processing (see below). Fertilize the remaining culture and mechanically remove fertilization envelopes prior to setting the culture. Synchronicity is evaluated microscopically and samples of all replica cultures should be cultured to ensure proper development through gastrulation to the larval stage.

2. Generally, the cultures will be at roughly 10% volume-volume in ASW. At desired times, transfer 2 × 50 mL of culture and isolate the eggs, zygotes or embryos by centrifugation at 250 × g (hand centrifuge or a clinical centrifuge) for 30 seconds at 15–16^o C.

3. Note the volume of packed eggs, zygotes or embryos and resuspend in 5 volumes of ice cold buffer G (pH 6.9 for unfertilized eggs or pH 7.4 for zygotes and embryos), and quickly wash 3X by gentle resuspension and centrifugation using 5 volumes of the appropriate ice cold buffer G each time.

4. Remove as much buffer G as possible after the final wash. Add an equal volume of Buffer G (of the appropriate pH), resuspend, and transfer the dense egg, zygote or embryo suspension to a pre-chilled glass homogenizer on ice.

5. Manually disrupt the eggs and zygotes by homogenization on ice using a pre-chilled Teflon pestle. Start with 5–10 strokes and monitor microscopically for lysis. Avoid introduction of bubbles during lysis. The goal is to obtain close to 100% lysis but to avoid over-homogenization so that the abundant yolk platelets (which are subsequently removed by centrifugation) are not disrupted.

6. Centrifuge the lysates at 4^o C for 15 minutes at 10,000g in a pre- chilled Sorvall SS34 rotor. If conducting a small-scale experiment, top speed in a microfuge for 20 minutes at 4^oC works well.

7. Transfer the soluble supernatant to a new tube and centrifuge again as a final clearing step to remove any remaining debris, yolk platelets or granules. We have found that this extra step is critical for subsequent steps.

8. Snap freeze small (100–250 μL) aliquots of the soluble cell free lysates in liquid nitrogen, and store at −80^oC. Save a few small aliquots in order to immediately quantify protein concentrations (blank against the pH specific buffer G + inhibitors) and to qualitatively assess sample via SDS-PAGE followed by immunoblot analysis for known proteins. Generally, starting with 2 × 50 mL of a 10% culture yields ~ 1 mL of 2–5 mg/mL soluble cell free lysate. This amount is adequate for multiple runs through the downstream protein analysis workflow (Figure 1). In fact, the samples sometimes require dilution prior to trypsin digestion.

Figure 1.

Workflow for the global phosphoproteomic and interactome profiling of eggs and at various times after fertilization. (A) The phosphopeptide separation and detection strategy uses hydrophilic interaction liquid chromatography (HILIC) combined with phosphopeptide enrichment via TiO₂. (B) Co-fractionating proteins are isolated by several methods including ion exchange chromatography and sucrose gradient fractionation are also resolved by HILIC after digestion. Analysis by tandem mass spectrometry (MS/MS) identifies proteins, quantified by spectral counts and ion intensities. Ortholog mapping to the human proteome is followed by integrative computational analyses.

B3. Downstream processing

1. Proteins prepared by this method can be directly trypsin digested (Guo et al, 2015) or split and subjected to multiple column chromatography methods to fractionate protein complexes (such as sucrose gradient fractionation, size exclusion and ion exchange chromatography; Wan et al, 2015) prior to digestion. Briefly, the isolated proteins are adjusted to 2 mg/mL, equilibrated in 50 mM Tris (pH 8.0), then reduced using 5 mM DTT for 1 hour and 10 mM iodoacetamide for 45 minutes in the dark at RT. The adjusted protein sample are then digested overnight using a 20:1 protein:trypsin ratio. Following digestion, desalt the peptides on C-18 columns (Toptip; Glygen) and lyophilize to dryness. If outsourcing the sample for Tof MS/MS analysis, the trypsin digestion step is usually included as part of the processing.

2. For phosphoproteomic analyses, phosphopeptides must be enriched. Total peptides are first fractionated on a TSK gel amide-80 column using an Agilent 1100 HPLC system to separate in the first dimension. A tenth of each of 20 fractions is directly injected into an HPLC-coupled mass spectrometer for total proteome analysis and the remaining 90% is enriched for phosphopeptides using TiO₂-coated magnetic beads prior to Tof MS/MS (Guo et al, 2015).

3. Once the peptide spectra are available, users have a variety of options for database searching and quantitation. We use MaxQuant (http://maxquant.org/) with a UniProt/Swiss Prot protein database FASTA file. Most methods allow users to set parameters for missed cleavage sites, carbamidomethylation of cysteine, N-terminal acetylation and oxidized methionine. Regardless, we recommend stringent 1% false discovery rate thresholds. The annotated sea urchin reference genome allowed for identification of roughly 90% of the peptides identified in recent proteomic studies (Guo et al, 2015; Wan et al, 2015).

4. Like many omics approaches, a large amount of in silico data will be obtained; the use of internal replicates (see above) greatly aids in the analysis of the data, especially in regard to quantitative site-specific phosphorylation or co-fractionation. A first step usually requires compilation of the echinoderm proteins to human orthologs. This enables the user to carry out analyses of predicted function and pathway enrichment (such as ROAST; Wu, Lim, Valliant, Asselin-Labat, Visvader & Smyth, 2010), including standard Gene Ontogeny (GO) and databases that have searchable pathways (cf http://baderlab.org/EM_genesets/). Several different platforms are available for visualizing enrichment pathways graphically; we use Enrichment Map, which is a plug-in for Cytoscape (Merico, Isserlin, Stueker, Emili & Bader, 2010). It is helpful to have a few predetermined signatures that are expected to appear in the analysis as an internal proof of concept and of course, to validate any candidates of interest through other means. Phosphorylation specific antibodies can be used to assess phosphorylation of individual candidate proteins over time after egg activation via immunoblotting while co-immunoprecipitation and protein affinity capture methods (Roux-Osovitz & Foltz, 2014) work well to interrogate protein-protein interaction candidates. Machine learning methods trained against existing databases such as CORUM (Ruepp, Waegele, Lechner, Brauner, Dunger-Kaltenbach & Fobo, 2010) are rapidly evolving to analyze protein interactomes with high precision (Havugimana, Hu & Emili, 2017).

IV. Nascent Protein Identification and Isolation

Identification of newly synthesized proteins is an important part of studying mechanisms of embryo development. The egg makes and stores many maternal proteins for use during embryogenesis so examining total protein in embryos needs to be interpreted with caution, as does morpholino antisense oligonucleotide (MASO) procedures when early in development the majority of protein is maternally derived. Further, identifying the dynamics and/or location of only the mRNA in the embryo is insufficient to deduce when and where cognate proteins accumulate. Many cases have reported the presence of mRNA in cells without protein (no translation of the mRNA, or rapid turnover of the protein (cf Voronina, Lopez, Juliano, Gustafson, Song & Extavour, 2008) or protein without detectable mRNA (long-lived protein; Song & Wessel, 2012). The following methodology is used to identify newly made protein at specific times in sea urchin development.

A. Identification of newly synthesized protein

A1. Methodologies to see specific protein synthesis

Specific protein identification is usually antibody based – using antibodies in immunoblots (Western blots) and in situ immunolabeling is the gold standard for endogenous protein localization on a candidate by candidate basis. Generating specific antibodies to echinoderm proteins is usually advantageous, but in some cases, commercially available antibodies to conserved proteins should also be considered. Making new antibodies is now possible from commercial sources by simply emailing the sequence of the immunogen and a check (cf Genescript, https://www.genscript.com/). Fortuitous cases also allow protein functional assays e.g. alkaline phosphatase activity, even in situ (Krugelis, 1947; Bishop, Bates & Brandhorst, 2002).

Sometimes specific protein localization can be mimicked by tagging the coding region of an mRNA and injecting the transcribed mRNA into the cell or early embryo. The investigator then identifies the exogenously expressed protein by fluorescence (e.g. GFP fusion protein) or by an epitope recognizable by a commercially available antibody (e.g. FLAG-tag fused to protein). The placement of the tag within the protein sequence is often important since one does not want it to interfere with the normal functionality or regulation of the protein. It is also an empirical science. Generally, it is best to place the tag at the C-terminus so that a signal indicates the encoded protein was at least translated in order to get to the tag. It is prudent to place such tags in the N-terminus as well as the C-terminus, minimally, to determine how much impact the tag has on the protein localization. In many cases this tagging approach closely replicates the endogenous protein synthesis (e.g. GFP-Nanos2 in S. purpuratus; Oulhen, Yoshida, Yajima, Song, Sakuma, Sakamoto Yamamoto & Wessel, 2013), but each case needs to be tested independently with an antibody to the endogenous protein.

A2. Methodologies to see general protein synthesis in the embryo

Detecting the synthesis of new macromolecules usually requires a labeled precursor molecule. Historically, the precursor molecules contained radioactive carbon, sulfur, or hydrogen. Identification of the synthesized molecule then required a detection system, usually film, or a film emulsion of silver halide crystals, in order to record the site of the radioactive signal. This approach was used extensively in echinoderms (e.g. Bedard & Brandhorst, 1983; 1986) to label newly synthesized proteins in early development.

Newer technologies now use non-radioactive precursors with easier and more varied detection methodology without sacrificing the ease of access for the substrate into the embryo. Here we document use of Click-iT technology for detecting general protein synthesis both in situ, and by Western blot analysis, and for isolation of newly synthesized protein.

A3. Nascent protein labeling in situ

Labeling of newly synthesized proteins is accomplished with the Click-iT protein synthesis assay kit (Life Technologies, Carlsbad, CA) with either OPP, O-propargyl-puromycin (C10457), or HPG, L-homopropargylglycine (C10428). OPP is a puromycin derivative, and as with its parent molecule, it causes premature chain termination perhaps from its structure that is similar to an aminoacylated tRNA with no ability for chain elongation. As a consequence, it is toxic to prokaryotic and eukaryotic cells when used for prolonged times (>30 min). HPG, on the other hand, is an amino acid analog of methionine that enables complete chain elongation and even functional proteins. Both products have an alkyne modification, will enter cells of the embryo, and are used to report sites of nascent protein synthesis (for sea urchin application see Oulhen et al, 2017). Detection of the incorporated Click-iT substrate is accomplished by fixing the cell and covalently labeling the nascent polypeptide with a fluorescent molecule containing the complementary azide chemistry that in the presence of copper ions, will covalently connect the labeling molecule with a fluorescence reporter (Figure 2).

Figure 2. Fluorescent reporters.

The Click-iT reaction has many variations in substrates and tagging methodologies. Shown here are possible precursor molecules that can be used and that are modified with an alkyne group. Once the precursor molecule is incorporated into the cell, and fixed in place, the detection molecule will spontaneously and non-enzymatically react with the alkyne group through its azide linkage to covalently label the site where the precursor molecule was fixed in place. Options for tagging molecules include fluorescence, if in situ labeling is the goal, or biotinylation, if isolation or biochemical detection in a blot is desired.

A routine protocol is as follows: Embryos are incubated for 30 minutes with the OPP used at 1:4000 (5 μM final), or HPG used at 1:2000 (25 μM final), then fixed for 15 minutes at room temperature (RT) with PFA 4% in ASW. Longer incubations with HPG are feasible, depending on the intent of the experiment, but not for OPP. After the PFA fixation, embryos are transferred to a 96 well plate in order to use smaller volumes of reagents. These wells are usable for up to 200 μl. The nascent proteins are then labeled according to the manufacturer’s instructions. Briefly, the embryos are washed one time with PBS - 0.5% Triton X-100 for 10 minutes at RT, three times with PBS - 0.5% Tween 20 (PBST) for 10 minutes each at RT, and two times with PBS only for 10 minutes each at RT.

A 1x OPP Click-iT reaction cocktail was made as follows (for 1 ml): 880 μl of Click-iT OPP reaction buffer (1x), 20 μl of Copper protectant, 2.5 μl of Alexa fluor azide, 100 μl of OPP Click-iT reaction buffer additive (1x). A 1x HPG Click-iT reaction cocktail was made as follows (for 1 ml): 860 μl of Click-iT HPG reaction buffer (1x), 40 μl of Copper protectant, 2.5 μl of Alexa fluor azide, 100 μl of HPG Click-iT reaction buffer additive (1x). The fixed embryos are incubated with the OPP or the HPG 1x Click-iT reaction cocktail, for 30 minutes at RT, in the dark. They are then washed one time with the corresponding wash buffers provided in the kits, and five times for 10 minutes each with PBST at RT. To label DNA, the embryos were incubated with the nuclear stain (diluted to 1:2000) provided in the kits, for one hour at RT in the dark and washed four times with PBST.

At the end of this protocol, the embryos were either washed overnight in PBST at 4°C before being imaged (this overnight wash is important to remove any potential background signal in the embryos before imaging) or incubated in the blocking buffer (PBST – sheep serum 4%) for an hour at RT, and then processed for immunolocalization for co-labeling purposes. Images are then best captured by confocal microscopy and fluorescence quantified using Metamorph or NIH ImageJ.

B. Identification and Isolation of newly synthesized proteins

If the goal of the experiment is to isolate and to identify the newly synthesized polypeptides, the investigator should then use an azide-conjugated biotinyl Click-iT reaction group instead of the fluorescence reporter for in situ nascent protein detection. Identification of nascent proteins by Western blotting: Seventy embryos (S. purpuratus blastula) are sufficient to detect a signal by blotting for many proteins of interest. After incubating the live embryos with OPP or HPG, the embryos are pelleted and lysed in 10 μl of loading buffer (5mM Tris pH 6.8, 1mM EDTA, 2% SDS, 20% sucrose, and Bromphenol blue). One microliter of 100 mM DTT was added to each sample before incubating them at 95^oC for 5 minutes. Each sample is loaded on a 4 to 12% gradient SDS-PAGE and transferred to the nitrocellulose membrane. This membrane is blocked with TBST- 1% BSA for one hour at RT. To Click-iT the biotin tag with the HPG or OPP now on the blot, the protein reaction buffer kit is used (C10276, Life Technologies, Carlsbad, CA).

The Click-iT reaction is made as follows: 500 μl of Click-iT reaction buffer containing 5 μl of Biotin Azide of a 4 mM stock (B10184, ThermoFisher Scientific, Waltham, MA), 300 μl of water, 50 μl of CuSO4, 50 μl of Click-iT reaction buffer additive 1 solution. The tube is vortexed for 5 seconds after addition of each reagent to the Click-iT reaction. Two minutes later, 100 μl of Click-iT reaction buffer additive 2 is added before vortexing the mix for 5 seconds. The nitrocellulose membrane is incubated with this 1 ml mix at room temperature for 20 minutes, on a shaker. The membrane is then washed 3 times with TBST and incubated overnight with streptavidin horseradish peroxidase (ab7403) diluted to 1:1000 in the blocking buffer. On the following day, after three additional washes with TBST, the bands are observed by imaging chemiluminescence. Controls for this experiment should include comparing the experimental embryos to sibling embryos that received no biotinylation reagent, to test for endogenous biotin in the cells.

Isolation of newly synthesized proteins: Following HPG or OPP incubation during the time frame of interest, the embryo is lysed with detergent, and the biotinylated tag (prepared as above for blotting) is reacted with the lysate. Following this reaction, the sample is dialyzed against PBS-T with 3,000 MW cut-off dialysis tubing to remove excess biotinylation reagent. The biotinylated nascent protein then is isolated by use of streptavidin. We find that streptavidin magnetic beads work best for isolations since the beads are not pelleted to the bottom of the tube, potentially trapping background materials as in traditional bead purification reactions. The amounts of beads to lysate must be determined empirically for each sample. The isolated beads are washed 3 times with PBST and then resuspended for PAGE gel (SDS-PAGE loading gel buffer) or sent for mass spectroscopy analysis (trypsin digestion buffer). Controls for this experiment should include comparing the experimental embryos to sibling embryos that received no biotinylation reagent, to test for endogenous biotin in the cells.