extfieldsofstudy: Medicine Chemistry

s2fieldsofstudy: Biology Engineering

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Metabolic Engineering of Bacterial Respiration: High vs. Low P/O and the Case of Zymomonas mobilis

Respiratory chain plays a pivotal role in the energy and redox balance of aerobic bacteria. By engineering respiration, it is possible to alter the efficiency of energy generation and intracellular redox state, and thus affect the key bioprocess parameters: cell yield, productivity and stress resistance. Here we summarize the current metabolic engineering and synthetic biology approaches to bacterial respiratory metabolism, with a special focus on the respiratory chain of the ethanologenic bacterium Zymomonas mobilis. Electron transport in Z. mobilis can serve as a model system of bacterial respiration with low oxidative phosphorylation efficiency. Its application for redox balancing and relevance for improvement of stress tolerance are analyzed.

INTRODUCTION
During the last few decades a vast body of evidence on the structure, function and regulation of microbial electron transport chains has accumulated, enabled by the rapid advancement of omics techniques and molecular cloning tools. Research has focussed on deciphering of particular electron pathways in several model microorganisms, establishing their physiological role, and understanding the principles of their regulation. Transcriptional regulation in Escherichia coli in response to variation of available electron acceptors perhaps is the best known success story (for reviews, see Spiro and Guest, 1991;Unden and Bongaerts, 1997;Green and Paget, 2004). Here we briefly discuss how this wealth of theoretical knowledge can be used for industrial applications in the present era of metabolic engineering and synthetic biology. Generally speaking, microbial aerobic electron transport chains represent a promising, yet at the same time, challenging target for rational metabolic engineering, since their activities transcend several key cellular processes, like the catabolic ATP yield, maintenance of intracellular redox balance, production of reactive oxygen species (ROS), and protection against oxidative stress. In the present minireview we attempted to systematize the strategies of respiratory chain engineering, based on their impact on the cellular energy and reducing equivalent balance. We did not review the biotechnological applications of bacterial anaerobic respiration, electron transport chains of extremophiles, as well as bioelectrochemistry-interaction of bacterial electron transport with electrodes. Instead of trying to embrace all aspects of microbial electron transport, we here focussed on a few current biotechnology workhorses, showing how the metabolic engineering of their respiration had yielded predictable (or sometimes, not so predictable) effects on their growth and production. Some emphasis was put on low energy-coupling electron transport and the bacterium Zymomonas mobilis as a representative of the energetically uncoupled respiration.

RESPIRATORY ENGINEERING TO IMPROVE FERMENTATIVE CATABOLISM
Microbial respiration represents a major sink of reducing equivalents and a source of energy (ATP) for the cellular metabolism. However, if the target compound of a bioprocess is produced by a redox-balanced fermentative pathway, then respiration, by interfering with the cellular redox and energy balance, causes a decrease of product yield, to mention Pasteur effect as a classical example (for a review, see Barnett and Entian, 2005). NADH supply of reactions, producing the desired target compound of fermentative catabolism, often represents a bottleneck (Zhao et al., 2017), especially under aerobic or microaerobic conditions (Bennett and San, 2017). Removing oxygen from the growth medium seems a straightforward approach for turning off the unwanted respiration and fueling more NADH into the fermentative catabolism. However, prolonged maintenance of a strict anoxic condition may be a challenging task. It may complicate, for example, long-term directed evolution experiments, like those aimed at engineering of the E. coli fermentative metabolism (Portnoy et al., 2008), where complete absence of oxygen is essential. On the other hand, many yeast species performing ethanol fermentation require limited presence of oxygen (Visser et al., 1990), often within a certain narrow concentration range for optimum ethanol yield (Acevedo et al., 2017). From the technical point of view, keeping low, steady, micro-aerobic oxygen levels in bioreactors (especially for larger volumes) may be difficult, due to the sensitivity of oxygen mass transfer coefficient to the agitation rate, insufficient responsiveness of the dissolved oxygen probe and spatial in homogeneities (Wu et al., 2015a).
Decreasing the activity of electron transport chain itself represents a viable alternative way to downregulate respiration, so that the cells would behave similar to that of anoxic or microaerobic environment even in the presence of oxygen excess (Bennett and San, 2017). Such a metabolic engineering strategy of limiting the use of oxygen by the cell itself would be useful also under some co-cultivation settings, e.g., when an end-product of fermentative catabolism of one microorganism species is used as a substrate for aerobic metabolism of the second, co-cultivated species. Decreasing of respiration may be achieved by respiratory inhibitors, keeping in mind that inhibitor effects may serve as a proof of principle, yet are hardly applicable to industrial bioprocesses. Thus, Acevedo et al. (2017), using rotenone, the inhibitor of the mitochondrial NADH dehydrogenase complex I, demonstrated an increase of aerobic ethanol yield from xylose in the yeast Scheffersomyces stipitis. Cyanide at submillimolar concentrations was shown to improve growth and ethanol yield in aerobic culture of ethanologenic bacterium Zymomonas mobilis (Kalnenieks et al., 2000).
Not surprisingly, construction and study of respiratory knockout mutants nowadays has become the dominating approach to the downregulation of microbial respiration, and has brought positive results. In E. coli, removal of all its terminal oxidases, followed by adaptive evolution, yielded strains able to produce lactate as a fermentation product from glucose and to undergo mixed-acid fermentation during aerobic growth (Portnoy et al., 2008). For Corynebacterium glutamicum, another important biotechnological producer, a respiratory chain mutant lacking both the cytochrome bd branch and the cytochrome bc 1aa 3 branch was constructed (Koch-Koerfges et al., 2013), and its ability of fermentative growth without aerobic respiration was demonstrated. The mutant retained only 2% of the wild type oxygen consumption rate and displayed fermentative catabolism with L-lactate as major and acetate and succinate as minor products. In aerobically grown Klebsiella pneumoniae knocking out of its NADH dehydrogenase complex I (nuo) improved 2,3-butanediol production . In the ethanol-producing, facultatively anaerobic bacterium Z. mobilis inactivation of its Type II respiratory NADH dehydrogenase (ndh) largely improved growth and ethanol yield under aerobic conditions (Kalnenieks et al., 2008;Hayashi et al., 2011Hayashi et al., , 2012see below).
Some metabolic engineering studies have attempted finetuning of the respiration rate in E. coli. In order to mimic microaerobic metabolism of E. coli under condition of excessive aeration, Zhu et al. (2011) manipulated respiration rate of cells by externally adding various amount of coenzyme Q 1 to a strain lacking the ubiCA genes necessary for quinone biosynthesis. They demonstrated that synthesis of fermentation products, like lactate and ethanol, varied inversely with the added amount of Q 1 . In the further study (Wu et al., 2015a), a nearly theoretical yield of lactate production was achieved under fully aerobic conditions by introducing a tunable pathway competing with biosynthesis of the precursors of coenzyme Q 8 . This regulatory strategy was named by authors "metabolic transistor" (Wu et al., 2015b), allowing to control a large flux (electron transport) by a small change in the availability of one of the electron transport components (Q 8 ). An approach somewhat similar to that of Zhu et al. (2011) has been applied to Lactococcus lactis, in which the activity of respiratory chain naturally depends on externally added hemin, the precursor of the cytochrome heme cofactors (Liu et al., 2017). The authors constructed a multiple dehydrogenase knock-out strain of L. lactis with its respiratory chain as the only remaining NADH sink, and showed how the external control of respiration rate by hemin affected its growth and the degree of reduction of the synthesized products (acetoin vs. butanediol).
Bacterial respiratory chains can be engineered to control NADH/NAD + ratio and reducing equivalent fluxes also under anaerobic conditions. For enhancing NADH-dependent fermentative production in some strict anaerobes, the activity of electron transport chain could be used to generate NADH at the expense of other reduced cofactors, e.g., ferredoxin. This approach recently was demonstrated in Clostridium thermocellum (Lo et al., 2017), and in the extreme thermophile Caldicellulosiruptor bescii (Williams-Rhaesa et al., 2018), in which conversion of pyruvate to acetyl-CoA proceeds largely via pyruvate:ferredoxin oxidoreductase. By overexpressing Rnf, the membrane bound energy-conserving ferredoxin:NAD + oxidoreductase, NADH flux to the NADH-dependent ethanol synthesis could be improved, leading to a substantial increase of ethanol production in both these bacteria, and a concomitant decrease of production of unwanted side-products.
Notably, some recent works on modification of respiratory NADH dehydrogenase activity in aerobic or facultatively anaerobic bacteria reported unexpected, NADH dehydrogenasedependent metabolic shifts under strictly anaerobic conditions, which at present seem difficult to explain. Under aerobic conditions, manipulating the activity of either the NADH dehydrogenase complex I (nuo) or the Type II NADH dehydrogenase (ndh) was shown to affect the intracellular NADH/NAD + ratio, and accordingly, the catabolic product spectrum in a fairly predictable manner, as reported for several bacteria, e.g., Bacillus subtilis (Gyan et al., 2006), E. coli (Liu Q. et al., 2014), K. pneumoniae , and Streptococcus agalactiae (Lencina et al., 2018). Yet, for example in C. glutamicum, overexpression of the ndh gene under oxygen-deprived conditions also substantially decreased the NADH/NAD + ratio, at the same time increasing the glucose consumption rate (Tsuge et al., 2015). Here "oxygen-deprived conditions" referred to <0.01 ppm oxygen, and importantly, no alternative terminal electron acceptors were present in the medium. Similarly, Hayashi et al. (2015) found a strong dependence of NADH/NAD + ratio on the Ndh activity in complemented Z. mobilis ndh mutant strains under strictly anaerobic conditions. No electron acceptors alternative to oxygen are known for this bacterium. Steinsiek et al. (2014) noted that inactivation of the NADH dehydrogenase complex I (nuo) in E. coli affected not only its aerobic metabolism, but induced significant metabolic response also under anaerobic conditions, and without alternative external electron acceptors-again, conditions where the respiratory chain should not be active. They observed genes coding for TCA cycle enzymes being upregulated, and higher amounts of succinate being synthesized by the mutant. At the same time, they noted a substantial shift of the ArcA phosphorylation pattern in the nuo mutant relative to the parent strain. Earlier Perrenoud and Sauer (2005) demonstrated that ArcA-dependent transcriptional regulation controls TCA cycle in E. coli not only under (micro)aerobic, but also under fully anaerobic conditions. Yun et al. (2005) reported higher lactate and formate yields in E. coli ndh mutant and ndh nuo double mutant under strictly anaerobic conditions. Apparently, more research is needed to understand these anaerobic effects and their possible relation to the global regulatory mechanisms linking respiratory chain to the central metabolism. At the same time, it is important to take them into account for practical respiratory engineering.

IMPROVING RESPIRATORY ENERGY-COUPLING
The classical work of Bauchop and Elsden (1960) provided evidence that on average one mole of ATP could fuel synthesis of slightly above 10 gdrywt of bacterial biomass. With a few exceptions, this value (Y ATP ) was found to be roughly similar for different microorganisms (Stouthamer, 1977). The apparent Y ATP is far from being a true "biological constant" for all growth conditions, since it depends on cellular maintenance energy requirements and energy-spilling reactions (Russell and Cook, 1995). Yet, primarily it is the variation of energy generation efficiency, but not that of Y ATP , that underlies the observed vast differences of microbial growth yields. Regarding aerobic catabolism, the efficiency of energy generation is determined by the stoichiometry of oxidative phosphorylation (Calhoun et al., 1993). The stoichiometry of oxidative phosphorylation (P/O) is the molar ratio between the generated ATP macroergic phosphate bonds and the oxygen atoms reduced to water by the respiratory chain (Hinkle, 2005). The mechanistic (maximum) P/O value for each particular case can be estimated from the known proton motive stoichiometries of the respiratory complexes and H + -ATP synthase, keeping in mind that the real values are lower due to proton leaks and slips in the energycoupling mechanisms. Typically, the bacterial electron transport chains are branched, and for electrons traveling from NADH via the respiratory complexes to oxygen, their contribution to energy coupling strongly depends on the particular pathway that the electrons take. For example, in E. coli electron transport chain (for a review, see Unden and Bongaerts, 1997) the stoichiometry of proton translocation per one transported electron (H + /e) may vary between 1 and 4 (accordingly, H + /O varies between 2 and 8). Assuming the proton/ATP stoichiometry of the H + -ATP synthase close to 3 (Kashket, 1982), the theoretical lower and upper limits for P/O in E. coli appear to be 0.67 and 2.67, respectively. In reality, E. coli and most other bacteria with branched respiratory chains tend to employ their energetically most efficient and less efficient electron transport branches simultaneously, resulting in a submaximal net energy coupling efficiency, which is hard to measure directly under in vivo condition. A metabolic network model with a growth ratedependent biomass composition, used for simulation of E. coli chemostat cultivations on several carbon substrates at different dilution rates, has yielded an estimate of P/O close to 1.5 (Taymaz-Nikerel et al., 2010). Thermodynamic analysis suggests that the submaximal P/O values, typically in the range of 1.0-1.5, indicate a tradeoff between metabolic energy conservation and energy dissipation, ultimately directed toward maximization of the microbial growth rate (Westerhoff et al., 1983;Molenaar et al., 2009;Werner et al., 2010).
From a practical point of view, increasing of P/O represents a metabolic engineering strategy for improving microbial biomass yield (Figure 1). For increase of P/O, the electron transport branches with low coupling efficiency have to be inactivated, and/or those with high efficiency have to be overexpressed. Biomass yield (and in some cases, e.g., in lactic acid bacteria, also multiple stress tolerance; Zotta et al., 2013) can thus be improved, which is desirable for cost-efficient biomass production of starter cultures, for the net productivity of bioconversions, and for the production of biomass-related compounds such as proteins, storage polymers and vitamins (Minohara et al., 2002;Brooijmans et al., 2007;Pedersen et al., 2012;Richhardt et al., 2013;Liu Q. et al., 2014).
Apart from biomass, high P/O ratio has been shown to push the synthesis of various products via pathways that consume (or at least, do not produce) ATP (de Kok et al., 2012). This takes place because the energetically efficient respiration minimizes the amount of carbon substrate that has to be oxidized to generate ATP for needs of growth and maintenance. Given that the carbon source is not limiting and its uptake is rapid enough, the high P/O may facilitate production of incompletely oxidized compounds, either target products or, sometimes, undesired inhibitory metabolites. This phenomenon is known as "overflow metabolism" (Neijssel and Tempest, 1976; Figure 1). Enhancing product synthesis by elevating P/O has been a strategy of quite a few metabolic engineering projects. In most cases, elevation of P/O is achieved by knocking out the energetically least efficient terminal oxidase (typically cytochrome bd), and thus redirecting electron flow to energetically more efficient terminal oxidase(s). Poly-γ-glutamic acid (γ-PGA), a multifunctional biopolymer with various applications, is an example of a product, for which ATP supply is essential for its biosynthesis. By elimination of the cytochrome bd branch in Bacillus licheniformis respiratory chain, Cai et al. (2018) reached an increase of γ-PGA yield by almost 20%. Efficient supply of ATP is critical also for production of riboflavin. In a fed-batch culture of riboflavin-producing B. subtilis a 30% higher riboflavin titer was accumulated by the cyd mutant compared to the control strain , apparently because of redirection of the electron flow to the proton-pumping aa 3 terminal oxidase. Similar approach was used for improving of N-acetylglucosamine production in B. subtilis  and lysine yield in C. glutamicum (Kabus et al., 2007). Liu Q. et al. (2014) demonstrated that inactivation of the cytochrome bd-II terminal oxidase and/or the type II (energy non-coupling) respiratory NADH dehydrogenase results in a several-fold increase of poly(3-hydroxybutyrate) accumulation in an E. coli recombinant producer strain.
However, under some natural conditions, and also in numerous bioprocess setups it may be preferable for bacteria to have respiration with low rather than high P/O.

INCREASING RESPIRATION RATE WHILE DECREASING ENERGY-COUPLING EFFICIENCY
Low P/O is preferable when oxidation of NAD(P)H is needed for a rapid regeneration of NAD(P) + and/or consumption of oxygen, without producing of extra ATP (Figure 1). Thus, the free-living nitrogen-fixing bacteria Azotobacter vinelandii demonstrate the utility of rapid, low-P/O respiration as one of possible means to protect their nitrogenase complex from dioxygen by keeping the intracellular oxygen concentration low (Poole and Hill, 1997). It was shown that the activity of the low energy-coupling cytochrome bd terminal oxidase (Kelly et al., 1990) in combination with an energy non-coupling NADH:UQ oxidoreductase (Bertsova et al., 2001) is critical for A. vinelandii ability to fix nitrogen while growing under conditions of air saturation. In bioprocesses, energetically uncoupled respiration is useful for avoiding production of unneeded biomass and overflow metabolites, while enabling conversion of reduced substrate into more oxidized product(s). The obligately aerobic alpha-proteobacterium Gluconbacter oxydans belonging to the family of acetic acid bacteria is a relevant example. Due to its numerous periplasmic membrane-bound dehydrogenases that incompletely oxidize sugars, sugar alcohols, and other compounds, G. oxydans has been used for decades as a biocatalyst for bioconversions (Deppenmeier et al., 2002). It oxidizes a variety of substrates and transfers the electrons via ubiquinone to the quinol oxidases bo 3 and bd. Although the energetically more efficient, proton-pumping bo 3 represents its major terminal oxidase activity (Richhardt et al., 2013), due to some reason the proton pumping stoichiometry of the G. oxydans respiratory chain is more than two-fold lower than that of aerobically cultivated E. coli bearing analogous major terminal oxidase.
Seeking to relate the overflow metabolism phenomenon in the model microorganisms E. coli and S. cerevisiae to their respiratory capacity, Vemuri et al. (2006aVemuri et al. ( , 2007 demonstrated that overexpression of plant non-coupling alternative terminal oxidases, and/or the energy non-coupling, soluble water-forming NADH oxidase from Streptococcus mutans reduces the overflow metabolism, decreases the biomass yield, and shifts catabolism toward more oxidized products. In E. coli overexpression of streptococcal NADH oxidase was shown to raise the TCA cycle flux and to elevate the NADPH/NADP + ratio (Holm et al., 2010). In combination with the deletion of arcA gene it dramatically reduced acetate accumulation and hence improved capacity of recombinant E. coli to overexpress heterologous proteins (Vemuri et al., 2006b). Expression of the soluble water-forming NADH oxidases (nox) of Streptococci or Lactococci, which do not contribute to energy coupling, has been widely applied in metabolic engineering projects for enhancing respiration, lowering its energy-coupling efficiency, and decreasing the intracellular NADH/NAD + ratio. Overexpression of noxE from L. lactis in Saccharomyces cerevisiae has been shown to shift catabolism toward more oxidized products, like acetaldehyde, acetate, and acetoin (Heux et al., 2006). NoxE overexpression prevents accumulation of reduced byproducts, such as glycerol and xylitol, yet improves ethanol yield during xylose fermentation by engineered S. cerevisiae , as well as improves oxidative and osmotic stress tolerance of S. cerevisiae aerobic batch culture on glucose (Shi et al., 2016). The same oxidase has been overexpressed in E. coli (Balagurunathan et al., 2018) and in L. lactis (Bongers et al., 2005) to increase acetaldehyde yield at the expense of more reduced products of sugar catabolism. Bacterial soluble NADH oxidases are recently used also for in vitro metabolic pathways, reconstituted with isolated enzymes, as the means for recycling NADH (Nowak et al., 2015;Taniguchi et al., 2017).
In a broader sense, the metabolic engineering strategy for uncoupling of energy metabolism is not confined merely to manipulating the electron transport modules. A decrease of the energy-coupling efficiency can be achieved also by targeting the proton-dependent ATPase in order to disrupt oxidative phosphorylation, or to stimulate ATP hydrolysis. Following this approach, one can achieve energetical uncoupling in combination with overflow metabolism. Indeed, several studies have reported a decrease of biomass yield, growth rate, and intracellular ATP/ADP ratio, an increase of glycolytic and respiratory rates, and a rise of acetate production (overflow) in ATPase-deficient E. coli mutants (Jensen and Michelsen, 1992;Noda et al., 2006). Noda et al. (2006) observed activation of the low energy-coupling modules in the electron transport chain of such mutant: the type II NADH dehydrogenase and cytochrome bd terminal oxidase were upregulated. The reason for that might be the need for rapid regeneration of NAD + to keep the rate of glycolysis high, while at the same time investing less into proton motive force generation, since it no more could drive ATP synthesis. A similar shift in respiratory chain composition and an onset of acetate overflow were reported by Schuhmacher et al. (2014) in a wild type E. coli, grown under phosphate limiting condition. Notably, these physiological effects of phosphate limitation, resembling the ATPase knock-out phenotype, correspond to earlier in vitro findings with membrane preparations (Turina et al., 2004;D'Alessandro et al., 2008); and reconstituted liposome systems (D'Alessandro et al., 2011), indicating that H + -ATPases of E. coli and Rhodobacter capsulatus at low phosphate and ADP concentrations decouple ATP hydrolysis from transmembrane proton translocation. Causey et al. (2003) employed the acetate overflow effect for metabolic engineering of an acetate-producing E. coli. By inactivating the H + -ATP synthase along with disruption of the TCA cycle and elimination of native fermentative pathways, they obtained a strain, which synthesized acetate as its major product, reaching 86% of the theoretical maximum yield. Apart from E. coli, a somewhat similar response to inactivation of ATP synthase has been documented in other biotechnologically important bacteria, like C. glutamicum and Bacillus subtillis (Santana et al., 1994;Koch-Koerfges et al., 2012).
Alternatively, instead of disrupting oxidative phosphorylation, one can decrease the energetic efficiency of metabolism just by stimulating futile hydrolysis of ATP. In their pioneering work (Koebmann et al., 2002a,b) overexpressed the F 1 part of H + -ATPase, which could be used as an universal module for metabolic engineering of intracellular ATP hydrolysis without interfering with any particular metabolic reaction. In E. coli and L. lactis overexpression of F 1 led to a decrease of growth rate and intracellular energy level, and (depending on the strain and culture growth phase) also stimulated glycolytic flux to a various degree, decoupling it from anabolism. Using this approach Liu et al. (2016) demonstrated that futile hydrolysis of ATP could enhance product synthesis via pathways that generate ATP. By overexpressing of F 1 in L. lactis strain engineered for acetoin production they managed to improve the product yield and productivity at the expense of biomass formation. As noted by Holm et al. (2010), the metabolic effects of overexpression of the F 1 part of H + -ATPase and those of the soluble water-forming NADH oxidase had much in common. In both cases for E. coli the result was an elevated glucose consumption and reduced growth rate and biomass yield, although the acetate overflow was seen only with F 1 , but not with Nox.
Finally, since majority of bacterial electron transport chains are branched (for a review, see Poole and Cook, 2000), in principle it should be possible to decrease P/O also by inactivation of the energetically efficient electron transport branch(es), while leaving the less efficient, low-coupling ones active. However, in several cases inactivation of the energetically efficient routes has resulted in a drop of the bulk activity of mutant respiratory chain, actually causing poor growth and an elevation of overflow metabolism. Reported examples are the E. coli strains with linear electron transport chains (Steinsiek et al., 2014), or aa 3 knock-out mutant of B. subtilis , bearing the least energetically efficient electron transport components, yet accumulating more acetate than the respective wild types. Hence, turning a natural bacterial respiratory chain with an average P/O value into a low-P/O and high-rate one often implies complex genetic modifications and fine-tuning, including knocking out the high-coupling modules and concerted overexpression of low-coupling respiratory chain constituents. Obviously, the best alternative to such complex genetic engineering might be employing bacterial respiratory chains with a naturally low P/O, whenever possible. L. lactis is one of such bacteria, bearing a low energy-coupling, inducible respiratory chain, suitable for oxidizing surplus NADH, while little affecting cellular ATP production (Liu et al., 2017). Notably, although NoxE from this bacterium has been expressed in several microorganisms to elevate their respiration rate (see above), its own inducible respiratory chain proved to be more efficient than NoxE for regeneration of its own intracellular NAD + pool and for its oxidative stress protection. The ethanol-producing alphaproteobacterium Z. mobilis is another prominent example of aerobic metabolism with a naturally uncoupled respiration.

Zymomonas mobilis: AN EXAMPLE OF UNCOUPLED ENERGY METABOLISM
Zymomonas mobilis is a facultatively anaerobic bacterium with a very rapid and efficient homoethanol fermentation pathway (Rogers et al., 1982), involving the Entner-Doudoroff (E-D) glycolysis, pyruvate decarboxylase, and two alcohol dehydrogenase isoenzymes (Sprenger, 1996). Under anaerobic or microaerobic conditions up to 98% of substrate carbon is incorporated in ethanol. Metabolic engineering of Z. mobilis has largely focussed on biofuel production from variety of renewable substrates, as well as on increasing its stress-resistance (Zhang et al., 1995;Mohagheghi et al., 2015;Yang et al., 2016;Charoensuk et al., 2017; for recent reviews, see Wang et al., 2018;Xia et al., 2019). Fermentative catabolism in Z. mobilis is loosely linked to its anabolism and growth, and proceeds with a high rate, both during the exponential growth phase, or under non-or slowlygrowing conditions (Belaich and Senez, 1965). With respect to its metabolism, Z. mobilis somewhat resembles lactic acid bacteria (Sauer et al., 2017), also having high carbon substrate uptake rates with low biomass formation in combination with simple central metabolic pathways, leading to a limited number of metabolites. However, the aerobic energy metabolism in Z. mobilis seems to stand further away from the "bacterial mainstream, " than that of lactic acid bacteria.
In contrast to the inducible electron transport of lactic acid bacteria, which requires hemin to be present in growth media (Brooijmans et al., 2007;Koebmann et al., 2008), Z. mobilis bears a constitutive respiratory chain. It consists of the Type II NADH dehydrogenase (ndh), coenzyme Q 10 , and the cytochrome bd terminal oxidase (cydAB) as the established major electron carriers (Figure 2), together with respiratory FAD-dependent D-lactate dehydrogenase (Ldh), PQQ-dependent glucose dehydrogenase (Gdh), a putative cytochrome c peroxidase (PerC), and some other minor or still unidentified constituents (Strohdeicher et al., 1990;Kalnenieks et al., 1998;Sootsuwan et al., 2008). The cytochrome bd seems to be the only terminal oxidase, which is encoded in the Z. mobilis genome (Seo et al., 2005;Yang et al., 2009), and has been detected spetroscopically (Kalnenieks et al., 1998;Sootsuwan et al., 2008). Z. mobilis, like G. oxydans (Hanke et al., 2012) and some Acetobacter species, possesses a "dead-end" cytochrome bc 1 branch, apparently lacking a terminal cytochrome c oxidase, hence its function in electron transport remains obscure. The cytochrome c peroxidase seems more likely to function as a quinol peroxidase (Balodite et al., 2014), than to terminate the bc 1 electron transport branch, as suggested by Charoensuk et al. (2011). Nevertheless, Z. mobilis mutant strain with inactivated cytochrome b subunit of the bc 1 complex shows a distinct respiratory phenotype (Strazdina et al., 2012), best seen in mutant membrane preparations as an increase of antimycinresistance of oxygen consumption and alteration of cytochrome bd reduction kinetics with NADH. In G. oxydans its bc 1 -deficient strain shows a growth defect under acidic conditions, while in the wild type the qcrABC genes are upregulated under oxygen limitation (Hanke et al., 2012). Further research might reveal novel roles for the bc 1 complex in these alpha-proteobacteria and perhaps, put forward new ideas for their respiratory metabolic engineering.
For some reasons, the energetic efficiency of Zymomonas respiration in aerated culture is significantly lower than reported for other bacteria, e.g., L. lactis, with a similar dominating electron transport pathway from Ndh to CydAB (Brooijmans et al., 2007;Rutkis et al., 2014Rutkis et al., , 2016, which makes it quite unique among the producer microorganisms. Unlike such prominent facultatively anaerobic workhorses of biotechnology as E. coli and B. subtilis, aeration does not improve Z. mobilis biomass yield (Belaich and Senez, 1965; for a review, see Kalnenieks, 2006). Study of respiratory knock-out mutants (Strazdina et al., 2012;Rutkis et al., 2014) did not indicate presence of any putative low energy-coupling electron transport branch, to which one might ascribe the overall low energetic efficiency. Oxidative phosphorylation has been reported in non-growing cells and cell membrane vesicles, and cells are capable of ATP synthesis with artificially induced proton gradient (Kalnenieks et al., 1993). Yet, the proton-dependent ATPase in Z. mobilis under aerobic growth conditions operates in the direction of ATP hydrolysis . This is possibly because of some intrinsic deficiency of proton motive force maintenance, or due to an unidentified proton leakage pathway in growing cells.
Being an active sink of reducing equivalents, respiration lowers the ethanol yield, since it withdraws NADH from the alcohol dehydrogenase reaction. Hence, respiration causes accumulation of acetaldehyde, the inhibitory metabolic precursor of ethanol, which contributes to the poor aerobic growth of this bacterium, at the same time being a valuable target product of Z. mobilis aerobic bioprocess (Wecker and Zall, 1987). From the practical point of view, respiratory chain of Z. mobilis is well-suited for oxidation of NADH (and NADPH, since its Ndh has a good affinity for both cofactors) without stimulation of biomass growth. Aerobic acetaldehyde production by Z. mobilis so far is the best reported application of its respiratory potential. Recently, acetaldehyde-producing strains were developed, that reach 70% of the theoretical maximum acetaldehyde yield (Kalnenieks et al., 2019), as well as strains with the acetaldehyde-generating pyruvate decarboxylase reaction relocated to periplasm, in order to mitigate toxic effect of acetaldehyde on cell interior .
It is not obvious, what are the physiological function(s) of Z. mobilis respiration in nature. Possibly, the respiratory chain might be accelerating glucose consumption by lowering the intracellular NADH/NAD + ratio (Rutkis et al., 2016), like it has been shown previously for C. glutamicum (Tsuge et al., 2015). Rutkis et al. demonstrated that an active respiratory chain is advantageous for reaching higher initial glucose uptake rate in non-growing cells, thus improving their survival, when glucose is available in small amounts and for restricted periods of time (most likely, that is what happens during the natural life cycle of this bacterium).
Respiratory chain apparently plays some role in the protection of Z. mobilis against oxic, high temperature, inhibitor and salt stresses. Cytochrome c peroxidase (perC) knock-out mutant of the thremotolerant strain TISTR 548 shows impaired growth at elevated temperature (37 and 39 • C) (Charoensuk et al., 2011). Hypersensitivity to hydrogen peroxide (although, not as pronounced as in catalase-deficient strain) is another phenotypic trait of Z. mobilis perC mutants (Charoensuk et al., 2011;Balodite et al., 2014). In their recent work with TISTR 548, Charoensuk et al. (2017) presented results of transposon mutagenesis, followed by the mutant screening for the loss of thermotolerance. They identified 26 knock-out mutations, deleterious for thermotolerance, most of which were related to biosynthesis and maintenance of cell membranes and envelope. Yet, among the mutated genes there was the flavoprotein WrbA, a new type (type IV) of NAD(P)H:quinone oxidoreductase (Patridge and Ferry, 2006), thought to function as a protector against oxidative stress. Martien et al. (2019) noted upregulation of all respiratory chain components following exposure of a strictly anaerobic culture of Z.mobilis Zm4 to vigorous aeration, and suggested that the activity of PerC and cytochrome bd limits the toxic effects of oxygen. Interestingly, Ndh activity was shown to confer advantage for Z. mobilis aerobic growth and flocculation on minimal media (Jones-Burrage et al., 2019). Upregulation of ndh and some other oxidoreductase genes were observed under lignocellulose inhibitor stress (Chang et al., 2018). Hayashi et al. (2015) demonstrated positive impact of Ndh activity on Z. mobilis salt stress resistance. Likewise, activity of respiratory chain is positively correlated with acetic acid tolerance in Acetobacter pasteurianus (Wu et al., 2017), with multiple stress tolerance of Lactobacillus plantarum (Zotta et al., 2013), and with resistance of E. coli to critically high growth temperatures (Murata et al., 2018). Details of the mechanisms behind these general stress-protective effects, however, largely remain obscure and require further study.
Paradoxically, turning off Z. mobilis respiration also appears to be beneficial for its aerobic growth. The Z. mobilis ndh knock-out mutant shows improved aerobic growth and ethanol synthesis on rich growth medium (Kalnenieks et al., 2008;Hayashi et al., 2011Hayashi et al., , 2012, which partly can be explained by the decrease of acetaldehyde accumulation. At the same time, inactivation of the ndh gene elicits an adaptive response at the level of the electron transport chain and several oxidative stress protection systems (Strazdina et al., 2012). In particular, the mutant bears higher activity of respiratory D-lactate dehydrogenase (Ldh). Hayashi et al. (2011Hayashi et al. ( , 2012 analyzed a set of spontaneous Z. mobilis respiratory mutants with Ndh-deficiency, and observed their improved aerobic growth and ethanol synthesis at elevated temperature (39 • C). Recently Strazdina et al. (2018) demonstrated that the elevated Ldh activity of the ndh strain is crucial for its temperature-resistance. Inactivation of ldh against the Ndh-deficient background renders the non-respiring double mutant more temperature sensitive than the ndh strain, bringing it back to the level of the wild type. The protective function of the respiratory Ldh at elevated temperature needs further examination, in order to establish what specific electron pathway(s) starting from Ldh might be involved in Z. mobilis thermotolerance.

CONCLUSIONS AND OUTLOOK
Engineering of respiratory chain represents a potent lever to modify metabolism of biotechnological producer microorganisms. Rational metabolic engineering of culture growth and product synthesis, based on targeted modification of bacterial respiratory chain branches has been proven successful in numerous cases. At the same time, engineering of electron pathways can elicit complex responses, particularly concerning physiology of stress resistance, which may be difficult to explain within the present picture of electron transport. Modification of respiratory chain components can cause unpredicted effects on growth, catabolism and intracellular redox balance. Some effects, observed under strictly anaerobic conditions, possibly indicate gaps in our understanding of anaerobic electron transport pathways. Apparently, in order to uncover the full potential of respiratory metabolic engineering, more basic research on the structure and regulatory interactions between electron transport and the rest of microbial metabolism is needed. For broadening our knowledge of the alternative physiological roles of respiration, bacteria like Z. mobilis, which do not use their respiration for energy generation, might be helpful as experimental model systems.

AUTHOR CONTRIBUTIONS
All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine

text:
CD22 blockade restores homeostatic microglial phagocytosis in aging brains

Microglia maintain homeostasis in the central nervous system (CNS) through phagocytic clearance of protein aggregates and cellular debris. This function deteriorates during aging and neurodegenerative disease, concomitant with cognitive decline. However, the mechanisms of impaired microglial homeostatic function and the cognitive effects of restoring this function remain unknown. We combined CRISPR-Cas9 knockout screens with RNA-seq to discover age-related genetic modifiers of microglial phagocytosis. These screens identified CD22, a canonical B-cell receptor, as a negative regulator of phagocytosis that is upregulated on aged microglia. CD22 mediates the anti-phagocytic effect of α2–6-linked sialic acid, and inhibition of CD22 promotes the clearance of myelin debris, amyloid-β oligomers, and α-synuclein fibrils in vivo. Strikingly, long-term CNS-delivery of a CD22 function-blocking antibody reprograms microglia towards a homeostatic transcriptional state and improves cognitive function in aged mice. These findings elucidate a mechanism of age-related microglial impairment and a strategy to restore homeostasis in the aging brain.

and 3) are differentially expressed with age. To do so, we used CRISPR-Cas9 knockout screens 18,19 to discover genetic modifiers of phagocytosis in the microglia-derived cell line, BV2. Given cell number limitations 20,21 , primary microglia screens are not currently feasible. Although BV2 cells do not accurately recapitulate all features of microglia in vivo 22 , the two cell types express similar phagocytic machinery (Extended Data Fig. 1a).
Therefore, genetic modifiers of phagocytosis in BV2 cells are suitable candidates for subsequent validation in vivo. To identify genes that might be druggable, we used libraries of single-guide RNAs (sgRNAs) targeting 954 genes coding for transmembrane proteins and 2,015 genes coding for known drug targets, kinases, and phosphatases, with 10 sgRNAs per gene along with ~3,000 negative control sgRNAs.
Cas9-expressing BV2 cells were infected with these libraries and selected for stable sgRNA expression to obtain a pool of single-knockout cells (Fig. 1a). We incubated the cells with pH-sensitive fluorescent latex particles and sorted phagocytic and non-phagocytic populations by fluorescence-activated cell sorting (FACS) (Extended Data Fig. 1b, c). Next, we sequenced genomic DNA from each population, analyzed the sgRNA distributions between populations (Supplemental Table 1), and estimated the effect size and P-value of each gene knockout using casTLE 23 . These screens identified 286 significant hits, including multiple known negative and positive regulators of phagocytosis (Supplementary Table 2; Extended Data Fig. 1d, e, f, g, h).
To filter this list for genes that might contribute to age-related phagocytic decline, we performed RNA-seq on hippocampal microglia from young and aged mice and compared hits from our BV2 screen to genes differentially expressed with age in primary microglia. Within the subset of genes targeted by our CRISPR-Cas9 screen, we searched for ageupregulated microglial genes whose corresponding BV2 knockout promotes phagocytosis, or age-downregulated microglial genes whose corresponding BV2 knockout inhibits phagocytosis (Fig. 1a). These genes might have a functional role in age-related microglial impairment, either through increased inhibition or decreased activation of phagocytosis. Strikingly, CD22, a sialic acid-binding immunoglobulin-like lectin (Siglec) typically expressed on B-cells 24 , was the only gene that fit these criteria (Fig. 1b). Specifically, CD22 is significantly upregulated in aged microglia (Fig. 1b, c), corroborating previously published datasets 16,17 , and knocking it out in BV2 cells promotes phagocytosis ( Fig. 1d; Extended Data Fig. 1i, j, k; Supplementary Video 1, 2). CD22 negatively regulates B-cell receptor (BCR) signaling 25 , and has been reported to modulate leukocyte phagocytosis in fish 26 . However, its role in the regulation of mammalian phagocytosis is completely unknown.
Other Siglecs have been shown to regulate microglial phagocytosis 27 ,including suppression of amyloid-beta (Aβ) clearance by CD33 in Alzheimer's disease (AD) 28 . Based on our RNA-seq analysis and flow cytometry profiling of primary microglia, CD22 is the only mouse Siglec that is significantly upregulated with age in the hippocampus (Supplementary Table 3; Extended Data Fig. 2). Using a quantitative flow cytometry-based assay (Extended Data Fig. 3a), we determined that aged microglia display approximately three times more CD22 surface molecules than young microglia, comparable to the number of molecules on immature B-cells (Extended Data Fig. 3b). To assess the regional distribution of microglial CD22 expression, we performed multiplexed fluorescence RNA in situ hybridization (RNAscope) on five brain regions from young and aged mice. We probed for CD22 as well as Tmem119, a microglia specific marker 29 . Whereas CD22+Tmem119+ microglia were almost completely absent in the young brain, the aged brain contained a large proportion of these cells in every region that we assessed (Fig. 1f, g), particularly the thalamus and cerebellum. We did not observe CD22+ puncta outside of Tmem119+ microglia, corroborating previously published RNA-seq datasets 30 that show CD22 is expressed exclusively by microglia in the mouse CNS (Extended Data Fig. 3c, e, f).
CD22 mediates the anti-phagocytic effect of α2-6-linked sialic acid CD22 is canonically expressed on B-cells, where it negatively regulates BCR signaling by binding sialic acid and recruiting SHP-1 or SHIP-1 via immunoreceptor tyrosine-based inhibitory motifs (ITIMs) 31 . To search for possible signaling partners of CD22 on microglia, we re-analyzed our initial CRISPR-Cas9 screen for hits related to CD22 function. Surprisingly, CMAS, a key enzyme in sialic acid synthesis, and PTPN6, which codes for SHP-1, were among the most significant hits (Fig. 2a). Time-lapse microscopy confirmed that knocking out CMAS or PTPN6, or removal of sialic acid via treatment with sialidase or 3F ax -Neu5Ac, a sialic acid biosynthesis inhibitor, robustly promotes phagocytosis (Fig. 2b, c; Extended Data Fig. 4a, b, c, d, e), phenocopying CD22 ablation. However, genetic or pharmacological inhibition of both CD22 and sialic acid simultaneously did not produce an additive phagocytic effect ( Fig. 2d; Extended Data Fig. 4f, g), suggesting that sialic acid is involved in CD22-mediated inhibition of phagocytosis.
To assess inhibitory signaling downstream of CD22, we measured basal SHP-1 activity in BV2 cells and found that CD22 KO cells express a reduced ratio of active phosphorylated SHP-1 to total SHP-1 protein, comparable to CMAS KO cells (Fig. 2h). Furthermore, treatment with a CD22 ligand-blocking antibody (Extended Data Fig. 4i, j) decreased SHP-1 activity in a dose-dependent manner (Extended Data Fig. 4k). Taken together, these data support a model in which CD22 binds an α2-6-sialylated ligand and promotes inhibitory SHP-1 signaling to restrain the phagocytic capacity of microglia.

CD22 inhibition promotes microglial phagocytosis in vivo
To assess the functional significance of age-related microglial CD22 upregulation in vivo, we stereotactically co-injected myelin debris, a phagocytic substrate that accumulates in the aging brain, with a CD22 blocking antibody or an IgG isotype control antibody into opposite hemispheres of the same aged mouse (Fig. 3a). We labeled the myelin debris with a constitutively fluorescent dye to assess clearance and with a pH-sensitive dye to confirm engulfment and lysosomal trafficking within Iba1+ cells (Extended Data Fig. 5a, b).
After 48 hours, anti-CD22 treatment robustly promoted clearance of myelin debris compared to IgG (Fig. 3b, c). Because microglia appear to be the only cell type in the mouse CNS that express CD22 (Extended Data Fig. 3c), this effect is likely microglia-specific. Importantly, CD22 blockade did not influence the recruitment of microglia to the injection site relative to IgG (Extended Data Fig. 5c) or a stab wound control (Extended Data Fig. 5d, e), and IgG itself did not influence myelin clearance (Extended Data Fig. 5f). Nearly all Iba1+ cells at the injection site were Tmem119+ regardless of treatment (Extended Data Fig. 5g, h), suggesting that the pro-phagocytic effect of CD22 blockade is mainly mediated by resident microglia and not infiltrating peripheral macrophages 29 . We confirmed the pro-phagocytic effect of anti-CD22 treatment ex vivo using freshly isolated microglia from aged mice and pH-sensitive fluorescent latex particles (Fig. 3d). Next, we injected labeled myelin debris into the brains of aged CD22−/− mice or age-matched wild-type mice (Fig. 3e). We observed comparable clearance of myelin debris with genetic ablation of CD22 as with antibody-based blockade of CD22 (Fig. 3f). Furthermore, CD22 blockade ( Fig. 3g) or genetic ablation (Extended Data Fig. 5i) had no significant effect in the young mouse CNS, where myelin debris is efficiently cleared at baseline and CD22 is minimally expressed.
Next, we evaluated the effect of anti-CD22 treatment on microglial phagocytosis of oligomeric Aβ, a particularly neurotoxic protein aggregate that accumulates in the AD brain 34 . Compared to IgG, anti-CD22 treatment efficiently cleared injected Aβ oligomers in vivo ( Fig. 3h; Extended Data Fig. 5j, k, l). Interestingly, a larger percentage of residual Aβ in anti-CD22 treated hemispheres was contained in acidified lysosomes (Fig. 3i), suggesting that CD22 blockade promotes degradation of engulfed debris. In an analogous in vivo phagocytosis assay, we found that anti-CD22 treatment promotes the clearance of extracellular α-synuclein fibrils (Extended Data Fig. 5m, n, o), a pathological hallmark of Parkinson's disease. Taken together, these data suggest that CD22 is a broad negative regulator of microglial phagocytosis in the aged CNS.

Long-term CD22 blockade restores microglial homeostasis and improves cognitive function in aged mice
Aging and disease overwhelm the homeostatic function of microglia, leading to a distinctive transcriptional state 35 characterized by the downregulation of resting microglial genes and the upregulation of activated microglial genes. To assess the transcriptional effects of CD22 blockade, we implanted aged mice with osmotic pumps to continuously infuse a CD22 blocking antibody or an IgG control antibody directly into the cerebrospinal fluid for one month (Fig. 4a). As opposed to systemic antibody administration or CD22−/− mice, this intervention enabled CNS-specific targeting of CD22 without B-cell infiltration into the brain (Extended Data Fig. 6a) or significant leakage into the periphery (Extended Data Fig.  6b). We labeled the respective antibodies with a fluorescent dye in order to assess drug distribution within the brain parenchyma (Extended Data Fig. 6c) and target engagement with CD22 on microglia (Extended Data Fig. 6d).
Following one month of continuous infusion, we performed RNA-seq on purified microglia from the hemi-brains of these mice contralateral to the cannulation site to minimize injuryinduced confounding factors. We found very few transcriptional changes between microglia from untreated and IgG-infused mice (Extended Data Fig. 7a). In contrast, we found 315 differentially expressed genes between IgG and anti-CD22 treated microglia meeting a false discovery rate (FDR) cutoff of 10% (Supplementary Table 4; Extended Data Fig. 7b). We compared these genes to those that are differentially expressed between microglia from young and aged mice 36 , and between microglia from wild-type and end-stage 5xFAD ADmodel mice 37 (Fig. 4b, c). Remarkably, anti-CD22 treatment promoted the upregulation of homeostatic microglial genes (e.g. Tgfbr1, P2ry13, Sall1, Il10ra, Mef2a;  Fig. 8c). Of note, CD22 blockade abrogated CCL3 secretion in the presence of oligomeric Aβ, but had no effect on basal levels.
To determine the effects of CD22 inhibition on age-related cognitive dysfunction, we assessed hippocampal-dependent learning and memory performance in aged WT and CD22−/− mice. Surprisingly, CD22−/− mice exhibited improved spatial memory in the forced-alternation Y-maze test (Fig. 4e) and improved associative memory in the contextual fear conditioning test (Fig. 4f). To more specifically assess the contribution of microglial CD22 to this behavioral phenotype, we performed the same tests in aged mice infused with IgG or anti-CD22 directly into the CNS via osmotic pump. Remarkably, blocking CD22 specifically in the CNS phenocopied the learning and memory improvements seen in CD22−/− mice (Fig. 4g, h), suggesting that microglial CD22 negatively and reversibly regulates cognitive function in the aging brain. Of note, we did not observe a significant cognitive improvement in aged mice treated systemically with anti-CD22 via intraperitoneal injection, suggesting that peripheral B-cells are unlikely to contribute to this effect (Extended Data Fig. 9a, b).
These hippocampal-dependent behavioral phenotypes were accompanied by an increase in the number of hippocampal Prox1+ dentate granule neurons expressing c-Fos, an immediately early gene induced by neuronal activation 39 (Fig. 4i, j). In addition, anti-CD22 treatment promoted a modest increase in phosphorylated-CREB (Extended Data Fig. 8d, e), a plasticity-related marker 40 , but did not alter hippocampal neurogenesis (Extended Data Fig. 8f). We observed no difference in C1q deposition (Extended Data Fig. 8g) or synaptic density, as assessed by the intensity of pre-and post-synaptic markers, synaptophysin and PSD-95 (Extended Data Fig. 8h, i).

Discussion
Here, we show that CD22 is a negative regulator of phagocytosis that is upregulated on aged microglia. CD22 mediates the anti-phagocytic effect of α2-6-linked sialic acid, and inhibition of CD22, either via antibody blockade or genetic ablation, promotes the clearance of myelin debris, Aβ oligomers, and α-synuclein fibrils in vivo. Long-term CD22 inhibition partially reverses the transcriptional signature of age-and disease-related microglia, increases markers of neuronal activation, and, ultimately, improves cognitive function in aged mice (Extended Data Fig. 10).
These findings raise important questions. First, what causes CD22 upregulation in the aging brain? Recent studies have highlighted the importance of environmental cues in shaping microglial identity 41,42,43 . Based on our recent scRNA-seq atlas of microglia during development 44 , CD22 is enriched in a subpopulation of postnatal day 7 microglia (Extended Data Fig. 3D) that engulf apoptotic oligodendrocytes and resemble disease-associated microglia of aging and neurodegenerating brains. Therefore, it is possible that CD22 is upregulated as a negative feedback mechanism to restrain excessive phagocytosis in response to overwhelming cellular debris in the developing brain. During aging, this protective developmental program might be inappropriately re-activated in response to increased myelin fragmentation and protein aggregation.
Correspondingly, how does CD22 inhibition improve cognitive function in aged mice? Our RNA-seq analysis revealed that anti-CD22 reduces the microglial expression of various neuromodulatory pro-inflammatory molecules. Age-related impairments in perivascular glymphatic 45 and meningeal lymphatic 46 systems have been shown to affect cognitive function, and CD22-regulated phagocytosis may cooperate with these clearance pathways to control debris accumulation and local cytokine concentration in the aging brain.
Similar to their dysfunction during normal aging, microglia in the AD brain display signs of impaired phagocytosis, and allelic variants in microglia-specific AD risk genes modify phagocytic function 28,47,48,49,50 . Several human RNA-seq datasets show that CD22 is upregulated in the AD brain 51 , enriched in subpopulations of microglia correlated with clinicopathological decline 52 , and, anomalously, expressed on human oligodendrocytes 53 . Furthermore, a variant in the INPP5D locus, which codes for the CD22 signaling partner SHIP-1, is associated with AD risk 54 . Determining whether CD22 function in the CNS is conserved between mice and humans will be an important step prior to translational studies.
In addition to AD, recent studies have found that CD22 is drastically upregulated in mouse models of amyotrophic lateral sclerosis 55 , a motor neuron disease, and Niemann Pick Type C 56 , a lysosomal storage disease. Future investigations of CD22 on microglia may illuminate therapeutic strategies to treat or prevent these neurodegenerative diseases via restoration of homeostasis in the CNS.

Animals
Aged C57BL/6 mice (18-24 months old) were obtained from the National Institute on Aging rodent colony. Young male C57BL/6 mice (2-4 months old) were obtained from Jackson Laboratories or Charles River Laboratories. CD22−/− mice were originally generated by L. Nitschke (University of Erlangen) and generously provided by James Paulson (Scripps Institute) and Romain Ballet and Eugene Butcher (Stanford University). All experiments used male mice, except aged WT vs CD22−/− behavioral experiments contained balanced groups of male and female mice. All animal care and procedures complied with the Animal Welfare Act and were in accordance with institutional guidelines and approved by the V.A. Palo Alto Committee on Animal Research and the institutional administrative panel of laboratory animal care at Stanford University.

Cell culture
BV2 cells were originally obtained from E. Blasi (Università di Modena e Reggio Emilia) and expanded in DMEM supplemented with 10% FBS, penicillin/streptomycin, and GlutaMAX (Thermo Fisher Scientific). HEK293T cells were cultured in DMEM supplemented with 10% FBS, penicillin/streptomycin, and GlutaMAX. All cells were maintained in a humidified incubator containing 5% CO 2 at 37 °C.

CRISPR-Cas9 Screen
The 10-sgRNA-per-gene CRISPR/Cas9 deletion library was synthesized, cloned, and infected into Cas9-expressing BV2 cells as previously described 18 . Briefly, ~12 million (for Membrane Proteins sub-library) or ~24 million (for Drug Targets, Kinases, Phosphatases sub-library) BV2 cells stably expressing EF1alpa-Cas9-BLAST were infected with the 10 guide/gene sgRNA sub-libraries (Addgene cat nos. 1000000122 and 1000000124) at an MOI < 1. Infected cells underwent puromycin selection (1.5 μg/mL) for 5 days after which point puromycin was removed and cells were resuspended in normal growth media without puromycin. After selection, sgRNA infection was confirmed by flow cytometry, which indicated >90% of cells expressed the mCherry reporter. Sufficient sgRNA library representation was confirmed by deep sequencing after selection. Cells were cultured and maintained at 1,000x coverage for one week. Phagocytic screening prey were prepared by labeling 3 μm amino-coated polystyrene particles (Polysciences) with CypHer5E pHdependent fluorescent dye (GE Life Sciences). For the membrane proteins screen, 12.5 million cells were sorted per replicate (>1000x coverage) with 55.2% in the non-phagocytic gate and 11.2% in the phagocytic gate. For the drug target, kinase, phosphatase screen, 24 million cells were sorted per replicate (>1000x coverage) with 53.1% in the non-phagocytic and 19.4% in the phagocytic gate. At the end of each screen, genomic DNA was extracted for all populations separately using a QIAGEN Blood Midi Kit. Deep sequencing of sgRNA sequences on an Illumina Nextseq was used to monitor library composition. Guide composition was analyzed and compared to the plasmid library and between conditions using casTLE (https://bitbucket.org/dmorgens/castle). Briefly, casTLE compares each set of gene-targeting guides to the negative controls, comprising non-targeting and non-genic ("safe-targeting") sgRNAs, which have been shown to more aptly control for on-target toxicity due to endonuclease-induced DNA damage. The enrichment of individual guides was calculated as the log ratio between phagocytic and non-phagocytic populations, and gene-level effects were calculated from ten guides targeting each gene. P values were then calculated by permutating the targeting guides as previously described 23 .

Primary Microglia Isolation
Primary mouse microglia were isolated as previously described 20 . Briefly, mice were transcardially perfused with ice-cold HBSS containing glucose and HEPES, and brains were removed into the same medium containing DNAse I. On ice, brains were minced with a razor blade and a single cell suspension was obtained by gentle Dounce homogenization. The suspension was filtered through a 100 micron strainer. Myelin was removed using magnetic myelin removal beads (Miltenyi Biotec). The remaining myelin-depleted cell suspension was stained with antibodies to distinguish microglia by flow cytometry. Previous studies have demonstrated the intense autofluorescent signal of microglia in the aging brain, likely due to the accumulation of the age-related pigment, lipofuscin 14 . In order to measure the expression of proteins on aged microglia by flow cytometry, we developed a gating scheme (Extended Data Figure 3F) to exclude autofluorescent cells (~10% of total microglia) from our analysis. Specifically, we excluded cells with high side-scatter signal which coincided with the autofluorescent population. Whenever possible, we used fluorophores with emission spectra in far red channels (e.g. APC-Cy7). Although this technique excludes an interesting age-specific population from our analyses, it enables confident cell-surface profiling of the majority of aged microglia by flow cytometry. For RNA-seq, all microglia (including autofluorescent cells) were collected.

RNA-sequencing and analysis
20,000 microglia were isolated from either hemibrains or micro-dissected hippocampi as described above, and sorted into RLT Plus buffer (Qiagen) containing beta-mercaptoethanol. RNA was extracted using a RNeasy Micro Plus kit (Qiagen) according the manufacturer's protocol. RNA integrity was assessed on a Bioanalyzer (Agilent), and high quality samples were used for library preparation. cDNA synthesis and amplification was performed using the SmartSeq v4 Ultra-low input kit (Takara), and libraries were tagmented, adaptor ligated, and indexed using the Nextera XT kit (Illumina). After normalization and pooling, libraries were sequenced on a Hiseq 4000 (Illumina) using paired-end 100bp reads. Libraries were sequenced to a depth of >30 million reads per sample. Raw sequencing files were demultiplexed with bcl2fastq, reads were aligned using STAR, the count matrix was generated using SummarizedExperiment, and differential expression analysis was performed using DESeq2 with standard settings.

Quantitative flow cytometry
Primary microglia from young and aged mice were isolated as described above. For the same mice, peripheral blood was collected with EDTA anticoagulant by terminal intracardial bleeding. Whole blood was mixed 1:1 with PBS and layered over Ficoll-Paque Plus (GE), spun at 400 x g for 30 minutes with no break, and the peripheral blood mononuclear fraction was collected. Microglia and peripheral blood cells were stained with immunophenotyping antibodies as well as PE-conjugated anti-CD22 (Ox97, Biolegend). Quantibrite-PE (BD) beads were resuspended in FACS buffer and analyzed on a BD LSRFortessa. Using the same laser powers and detector voltages, peripheral blood and microglia samples were analyzed for CD22 expression. A standard curve was constructed to correlate PE intensity with number of PE molecules per Quantibrite bead. Finally, CD22 molecule numbers on microglia and peripheral blood cells were calculated by interpolation.

qPCR
Microglia were isolated by FACS and RNA was isolated as described above. cDNA synthesis and amplification were performed with a SMART-seq V4 Ultra-Low Input RNA Kit (Clontech). Samples were diluted and mixed with SYBR green master mix before loading as technical triplicates for qPCR on a LightCycler 480 (Roche). ΔΔC T values normalized to β-actin were used to assess relative gene expression between samples. The following validated primer pairs for murine CD22 were used: 5'-CCA CTC CTC AGG CCA GAA ACT-3' (forward) and 5'-TGC CGA TGG TCT CTG GAC TG-3' (reverse).

In vitro phagocytosis assay
For in vitro phagocytosis assays, BV2 cells were split into a 24-well plate 24 hours prior to feeding at a density of 50,000 cells/well in serum free medium. Following specific treatments, wells were washed twice with PBS and fed pHrodo red-labeled 3 micron aminocoated polystyrene particles (Polysciences) at a ratio of 10 particles per cell. 4 phase and red fluorescent images per well were acquired every hour for 24 hours using the Incucyte S3 live cell analysis system (Essen Bioscience). For each time point, we calculated normalized phagocytosis using the following formula: red object area / phase confluence. To combine data from independent experiments, we calculated phagocytosis relative to control by setting phagocytosis at zero hours in the control condition to 0, and this value at 24 hours to 1. Technical triplicates for each experiment were averaged, and the average of three independent experiments was reported.

RNAScope in situ hybridization
RNA in situ hybridization was performed on fresh frozen brain tissue using the Multiplex Fluorescence v2 kit (Advanced Cell Diagnostics) according to the manufacturer's protocol. Probes for CD22 and Tmem119 were commercially available from the manufacturer.

Lentivirus production and infection
HEK293T cells were transfected with packaging plasmids and sgRNA-containing plasmids. Supernatant was harvested at 48 hours and 72 hours and concentrated with Lenti-X solution (Clontech). BV2 cells stably expressing Cas9 endonuclease under blasticidin (1 μg/mL) selection were spin infected with lentivirus containing sgRNA plasmids under puromycin selection. Puromycin selection (1.5 μg/mL) was started 48 hours after infection and maintained for 7-14 days. Antibiotic selection was subsequently removed for expansion and freezing. CMAS and CD22 KO BV2 cells were sub-cloned by single-cell sorting to obtain a monoclonal knockout population. Other single-knockout cell lines were assayed as a polyclonal population.

Sialidase Expression
Sialidase from Vibrio cholerae was prepared as previously described 57 . In brief, Escherichia coli C600 transfected with the pCVD364 vector were grown in 2xYT media supplemented with ampicillin (100 mg/mL) for 12 hours at 37 °C. Cells were collected by centrifugation at 4,500 x g for 10 minutes. The pellet was resuspended in osmotic shock buffer (20% sucrose, 1 mM EDTA, 30 mM Tris-HCl, pH 8), incubated for 10 min at RT, then spun at 13,000 x g for 10 minutes. The pellet was resuspended in ice cold ddH2O, incubated at 4 °C for 10 min, and then spun at 13,000 x g. The supernatant was concentrated using a tangential flow filtration system (Pall Corporation). The concentrate was loaded onto a HiTrap Q HP anion exchange column (GE Healthcare Life Sciences) and eluted using a gradient of NaCl in 20 mM Tris, pH 7.6. Fractions containing sialidase were assessed by SDS-PAGE and pooled. Endotoxins were removed using a high capacity endotoxin removal kit (Thermo Fisher Scientific).

Sialic Acid Inhibition
For sialidase treatment, cells were incubated with 400nM Vibrio cholerae sialidase for 1 hour at 37 °C in serum-free DMEM prior to phagocytosis. 3F ax -Neu5Ac (R&D) was used as previously described. Briefly, the inhibitor was used at a concentration of 32 μM serially diluted in DMSO and used at a final concentration of 0.1% DMSO in DMEM. 3F ax -Neu5Ac was added with serum treatments for 48 hours prior to phagocytosis assays, and replaced in the feeding medium.

Glycopolymer Decoration
Aminooxy glycan-conjugated polymers were based on a poly(methyl vinyl ketone) backbone coupled to a dipalmitoylphosphatidylethanolamine lipid anchor. Glycopolymers were conjugated at 70-85% of ketone sites to either Neu5Ac-α2-3-LacNAc or Neu5Ac-α2-6-LacNAc through oxime chemistry. Cells were decorated with aminooxy glycanconjugated polymers as previously described 32 . Briefly, cells were harvested and resuspended in PBS at 10 7 cells/mL. Labeling was carried out by incubating the cell suspension with 1 μM polymer for 30 minutes at room temperature with constant gentle agitation. Cells were washed with PBS and split into a 24-well plate at 50,000 cells/well for subsequent phagocytosis assays.

Western blot
Protein lysates were prepared by incubating cell pellets on ice in Pierce IP Lysis Buffer or RIPA Lysis Buffer with cOmplete protease inhibitor cocktail (Thermo Fisher Scientific) and spun at 13,000 x g for 10 minutes. The supernatant was collected and protein concentration was measured with the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). An aliquot containing 5-20ug of protein from each sample was mixed with 4X loading buffer (Thermo Fisher Scientific) and boiled for 5 minutes at 95 °C before being subjected to SDS-PAGE and transferred to a nitrocellulose membrane (Bio-Rad). The membrane was first blocked with Odyssey Blocking Buffer (LI-COR) and stained overnight at 4°C with primary antibodies at the designated concentrations: mouse anti-α-tubulin (1:10,000, T9026, Sigma), rabbit anti-pSHP-1 (1:5,000, D11G5, Cell Signaling), rabbit anti-Sall1 (1:1000, ab31526, Abcam). The membrane was washed, stained with IRDye conjugated secondary antibodies (1:15,000, LI-COR) and imaged on the Odyssey CLx (LI-COR). The membrane was stripped by incubation with ReBlot Plus Strong Antibody Stripping Solution (Millipore) for 15 minutes before blocking, staining with anti-SHP-1 (1:10,000, C14H6, Cell Signaling) then an IRDye conjugated secondary antibody, and reimaged as described. Images were analyzed for band intensities with the ImageStudio software (LI-COR).

Antibody-mediated inhibition assay
WT BV2 cells were incubated in serum-free DMEM containing varying concentrations of a monoclonal anti-CD22 antibody (Cy34, BioXCell) from 20ug/mL to 0ug/mL for 8 hours at 37 °C. The p-SHP-1:total SHP-1 ratio, indicative of signaling activity downstream of CD22, was determined by phospho-western blot as described above.

Ligand-binding assay
Recombinant mouse CD22-human Fc fusion protein (R&D) was pre-complexed to an AF647-conjugated anti-human IgG secondary antibody on ice for 30 minutes at equal concentrations (10 μg/mL). The complex was then incubated with various concentrations of a mouse monoclonal anti-CD22 antibody (Cy34, BioXCell) or a mouse IgG1 isotype control antibody (MOPC21, BioXCell) on ice for 30 minutes. Next, the antibody-treated complexes were incubated with WT or sialidase treated BV2 cells for 30 minutes on ice. The cells were washed and rCD22 binding was assessed by flow cytometry on a BD Accuri C6.

Antibody internalization assay
WT BV2 cells were split into a 24-well plate 24 hours prior to antibody treatment at a density of 50,000 cells/well. Cells were treated with rat anti-mouse CD16/CD32 Fc Blocker (2.4G2, BD) then incubated with the following antibodies at 1ug/mL: mouse IgG1 isotype control antibody (MOPC21, BioXCell), mouse anti-CD22 (Cy34, BioXCell) and rat anti-CD22 (OX-96, Bio-Rad). All antibodies were pre-labeled with Incucyte FabFluor-pH Red (Essen Bioscience) according to the manufacturer's protocol. 9 phase and red fluorescent images per well were acquired every hour for 24 hours using the Incucyte S3 live cell analysis system (Essen Bioscience). For each time point, we calculated antibody internalization using the following formula: red object area / phase confluence. Technical triplicates for each experiment were averaged and reported.

Immunohistochemistry
Mice were euthanized with 2.5% (v/v) Avertin and transcardially perfused with ice-cold HBSS containing glucose and HEPES, or with 4% PFA. Hemibrains were post-fixed in 4% PFA at 4 °C overnight before preservation in 30% sucrose in PBS. Hemibrains were sectioned into 40 micron coronal slices on a microtome and stored in cryoprotective solution at −20 °C. Free-floating sections were permeabilized, blocked, and stained overnight at 4°C with the following primary antibodies at the designated concentrations: goat anti-Iba1 . Sections were washed, stained with Alexa Fluorconjugated secondary antibodies (1:250), mounted and set under a coverslip before imaging on a confocal laser-scanning microscope (Zeiss LSM880). Thioflavin S staining was performed after secondary antibody staining. Briefly, sections were stained in a 0.1% (w/v) solution for 5 minutes, followed by 3 washes in 50% EtOH, and a 30 minute rehydration in diH2O prior to mounting.

Preparation of fluorescent phagocytic material
Purified CNS myelin was isolated as previously described 58 . Briefly, mice were euthanized and their brains homogenized. Crude brain homogenate was subjected to a series of sucrose gradients under ultracentrifugation followed by osmotic shocks in hypotonic buffers to remove myelin sheaths from severed axons. Aβ oligomers were prepared as previously described 59 . Briefly, Aβ 1-42 (Anaspec) was treated with hexafluoroisopropanol (HFIP) to achieve a monomeric solution. Oligomerization was induced by diluting a 5mM monomeric Aβ DMSO stock to 200μM in ice-cold PBS and incubated overnight at 4 degrees. Preformed α-synuclein fibrils were prepared as previously described 60 . Phagocytic material was conjugated to fluorescent dyes as previously described 21 . For double-labeled myelin, 50 mg/mL of myelin prep was incubated with equimolar concentrations of AlexaFluor555-NHS ester and CypHer5E-NHS ester in PBS with 0.1M sodium bicarbonate for 45 minutes at room temperature. Myelin was washed 4 times with PBS and centrifugation to remove free dye. For Aβ and α-synuclein, 100μL of the 200μM protein aggregates were incubated with 4μL CypHer5E-NHS ester in PBS with 0.1M sodium bicarbonate for 30 minutes at room temperature. Free dye was removed using BioSpin Micro P-6 desalting columns (BioRad).

In vivo phagocytosis assay
CypHer5E and AlexaFluor555 double-labeled myelin (25mg/mL), CypHer5E-labeled Aβ oligomers (1mg/mL), or CypHer5E-labeled alpha-synuclein fibrils (1mg/mL) were mixed with a mouse IgG1 isotype control antibody (MOPC21, BioXCell, 1mg/mL) or anti-CD22 (Cy34, BioXCell, 1mg/mL) and injected into opposite hemispheres of aged mice using a stereotaxic apparatus (Kopf instruments). Mice were anesthetized using isofluorane, their skulls were exposed and a hole was drilled at the injection site using aseptic technique. One microliter of the antibody/phagocytic target material mixture was injected at +/− 2mm lateral, 0mm anterior-posterior, and −1.5mm deep relative to the intersection of the coronal and sagittal suture (bregma) at a rate of 200nL/minute. The needle was left in place for an additional 3 minutes to allow for diffusion, then slowly withdrawn. Mice received postsurgical buprenorphine and baytril for pain and infection prevention, respectively. After 48 hours, mice were anesthetized and transcardially perfused with 4% PFA. The entire injection site was sectioned, stained, and imaged as described above, and 6-8 consecutive sections, spaced 40 μm apart, were quantified to assess total target clearance over the entire area of the injection site. Sections at the margins of the injection site without noticeable Iba1+ microgliosis (which takes >48 hours to resolve and is not influenced by treatment) were excluded.

Ex vivo microglial phagocytosis assay
Whole brain myelin-depleted single-cell suspensions were prepared as described above. Next, a rough estimation of microglia counts from each brain was determined by strictly gating a volume-designated aliquot of the suspensions by forward scatter/side scatter on a flow cytometer (BD Accuri C6). Each suspension was adjusted to contain equal microglia per unit volume. Then, single cell brain suspensions were treated accordingly for 1 hour in serum free DMEM-F12 at 37 °C with periodic agitation. Treatments were washed out and cells were resuspended in FACS buffer containing pHrodo-or CypHer5E-conjugated particles at a ratio of 50 particles per microglia. The cell-particle mixture was incubated for 1-2 hours at 37 °C with periodic agitation. Phagocytosis was stopped by transferring the suspensions to ice, where the cells were stained with antibodies to distinguish microglia. Microglial phagocytosis was assessed by flow cytometry by pre-gating live CD11b+CD45lo cells and assessing pH-dependent particle fluorescence within this population. The eating gate was determined using an unfed control, and confirmed by sorting an aliquot of the sample into non-eating and eating populations by FACS followed by microscopic assessment.

Osmotic pump intracerebroventricular delivery of anti-CD22
A monoclonal anti-CD22 antibody (Cy34, BioXCell, 1mg/mL) or mouse IgG1 isotype control antibody (MOPC21, BioXCell, 1mg/mL) were conjugated to AlexaFluor647 (ThermoFisher) using NHS chemistry at a dye:protein molar ratio of 4. Free dye was removed and buffer was exchanged with artificial CSF (Tocris) using 40kD MWCO Zeba spin desalting columns (ThermoFisher). The labeled antibodies were loaded into 200 microliter osmotic pumps (Alzet/Durect) with a 28-day infusion rate of 0.25mL per hour. An infusion rate was chosen to maintain a steady state concentration of 10 μg/mL, a dose shown to promote phagocytosis with this particular antibody ex vivo. Osmotic pumps were connected to a cannula (Brain infusion kit III, Alzet) inserted at +1mm lateral, −0.3mm anterior-posterior, and −3mm deep relative to bregma in order to target the right lateral ventricle. The pump was placed subcutaneously and mice received post-surgical buprenorphine and Baytril.

Quantification of antibody leakage into the periphery following intracerebroventricular administration.
1 mg of CD22 antibody was incubated with 30x molar ratio of Trans-Cyclooctene-NHS ester (Click Chemistry Tools) overnight at 4°C before being quenched with 1M Tris pH 8.0 and desalted (Zeba, 40K MWCO, Thermo). Mice were administered unlabeled or Trans-Cyclooctene (TCO)-labeled CD22 antibody via osmotic pump intracerebroventricularly as detailed above, or with equimolar amounts intravenously (retro-orbital) or intraperitoneally. Seven days later, blood was collected with EDTA anticoagulant by terminal intracardial bleeding. EDTA-plasma was isolated by centrifugation at 1,000g for 15 min at 4 °C before aliquoting, flash freezing, and storage at −80 °C. Protein concentrations of plasma samples and unlabeled and TCO-labeled CD22 antibodies were measured with a BCA Protein Assay Kit (Pierce). 75 μg of plasma was aliquoted, denatured in 1% SDS, and incubated for 90 minutes in the dark at room temperature with 6 μM SiR-tetrazine (Spirochrome) for subsequent click chemistry detection by in-gel fluorescence. 10x decreasing amounts of unlabeled and TCO-labeled CD22 antibody-from 1 μg to 0.1 ng-were processed similarly as a fluorescence standard. A 4x stock solution of NuPAGE LDS (Thermo) was added to each sample before heating at 95°C for 5 minutes. Proteins were briefly spun and separated by electrophoresis in 12% Bis-Tris polyacrylamide gels (Invitrogen). Gels were washed twice in distilled water for 15 minutes before SiR imaging in the 700-nm channel of an Odyssey CLx (LI-COR). To ensure accurate protein loading, gels were incubated with GelCode Blue Stain Reagent (Thermo) overnight before destaining in distilled water for at least 3 hours and imaged in the 800-nm channel of an Odyssey CLx (LI-COR). CD22 fluorescence signals were quantified in accompanying Image Studio™ software (LI-COR).

Behavioral assays
The forced alternation Y-maze was performed as previously described with minor alterations 61 . Briefly, the test consisted of a 5 minute training trial followed by a 5 minute retrieval trail, with a 1 hour inter-trial interval. For the training trial, one arm of the Y-maze was blocked off and mice were allowed to explore the two open arms. One hour later, the mouse was again placed in the Y-maze with all three arms open and a black and white pattern placed at the end of the novel arm. Between mice and trials, the maze was wiped with ethanol to remove odor cues. For analysis, video was analyzed by a blinded observer and both number of arm entries and time spent in each arm was quantified. Mice with less than two arm entries in the first minute of the retrieval trial were excluded from the analysis.
The fear-conditioning paradigm was performed as previously described 62 . Briefly, mice were trained to associate cage context or an audiovisual cue with an aversive stimulus (footshock). The test was administered over two days. On day 1, mice were placed in a cage and exposed to two periods of 30s of paired cue light and 1,000-Hz tone followed by a 2-s foot shock (0.6 mA), with a 180-s interval. On day 2, mice were subjected to two trials. In the first trial assessing contextual memory, mice were re-exposed to the same cage context, and freezing behavior was measured during minute 1-3 using a FreezeScan tracking system (Cleversys). In the second trial measuring cued memory, mice were placed in a novel context and exposed to the same cue light and tone from day 1 after 2 minutes of exploration. Freezing behavior was measured for 1-3 minutes following the cue. No significant differences in cued fear conditioning were observed between groups, consistent with previous rejuvenation studies from our lab 62 .

Assessment of the cognitive effects of systemically administered anti-CD22
Aged (18 m.o.) mice were intraperitoneally injected with 300μg of anti-CD22 (Cy34, BioXCell) or mouse IgG1 isotype control (MOPC21, BioXCell) twice weekly for one month, followed by cognitive assessment using the forced alternation Y-maze and contextual fear conditioning paradigms described above.

Gene Ontology Analysis
For gene ontology analysis of CRISPR-Cas9 screen hits, Panther was used to assess statistical overrepresentation of hits in the Reactome pathway database given a reference list containing all genes targeted by the sgRNA library. For analysis of RNA-seq data, Enrichr was used to assess enrichment of significantly up-or down-regulated genes in KEGG, BioCarta, WikiPathways, and Reactome databases. Pre-ranked gene set enrichment analysis was performed using GSEA software v3 (Broad Institute) with default settings and signed log-normalized p-values as a ranking metric.

Ex vivo microglia secretome profiling
Microglia from aged (20-24 m.o.) mice were isolated as described above, with minor modifications. Namely, following myelin depletion, cells were incubated with CD11bpositive enrichment beads (Miltenyi), isolated by magnetic selection using LS-columns, and plated at 30,000 cells / well of a 96-well plate. Microglia were maintained in serum-free defined medium as previously described 21 , and treated with either IgG (MOPC21, BioXCell, 20 μg/mL) or anti-CD22 (Cy34, BioXCell, 20 μg/mL) in the presence or absence of Aβ oligomers (5 μM) for 8 hours. Following incubation, supernatant was collected, spun down to remove cells, and flash frozen on dry ice. A bead-based immunoassay for mouse cytokines and chemokines (Eve Technologies) was used for protein detection.

Data Availability.
RNA-seq datasets have been deposited online in the Gene Expression Omnibus (GEO) under accession numbers GSE127542 and GSE127543. d, e, Results from CRISPR-Cas9 screen targeting 954 membrane proteins (d) or 2,015 drug targets, kinases, and phosphatases (e) in BV2 cells. Knockouts that promote phagocytosis (red) have a positive effect size and knockouts that inhibit phagocytosis (blue) have a negative effect size (screen performed in technical duplicate; dotted line, P=0.05, two-sided t-test). f, g, Distributions of negative control sgRNAs (gray) and Rab9-targeting (f) or CMAStargeting (g) sgRNAs (blue). Positive values indicate enrichment in the phagocytic fraction, and negative values indicate enrichment in the non-phagocytic fraction. h, Statistical overrepresentation test showing enrichment of Reactome pathway annotations within phagocytosis-promoting (red) and -inhibiting (blue) hits (Fisher's exact test). i, CD22 expression in WT (blue), CD22 KO (green), and isotype control stained (black) BV2 cells assessed by flow cytometry. j, Percent confluence of control (gray) and CD22 KO (green) BV2 cells during time-lapse microscopy phagocytosis assays (n=3, mean +/− s.e.m.). k, Number of beads ingested per cell were calculated in control and CD22 KO BV2 cells after 8 hours of phagocytosis. While CD22 KO cells display enhanced phagocytosis at a population level (n=3, *P<0.05, two-sided t-test, mean +/− s.e.m.), we observed no significant differences in the number of beads ingested per cell (two-way ANOVA). Red dot represents mean phagocytic index of the entire cell population. Data in b, c, i, j, k were replicated in at least 2 independent experiments.        i, Percent area of residual Aβ oligomers that were CypHer5E+, indicating localization to acidified lysosomes (n=8, **P<0.005, paired two-sided t-test). All data were replicated in at least 2 independent experiments. i, Representative images of dentate gyri stained for Prox1 (white) and c-Fos (red) from IgG and anti-CD22 treated mice. White arrows indicate cFos+Prox1+ active granule neurons. Scale bar = 100 microns. j, Total number of Prox1+c-Fos+ neurons in the dentate gyrus quantified over 5 tissue sections (n=7, *P<0.05, two-sided t-test; mean +/− s.e.m.). Data in a, g, h, i, j were replicated in at least 2 independent experiments.